Molecular Basis of Two Novel Dehalogenating Activities in Bacteria

Molecular basis of two novel dehalogenating activities in bacteria

Structure/function relationships in an evolutionary context

Foto’s omslag: Dinanda Lutikhedde, MFA

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The research presented in this work was performed in the protein crystallography group at the Laboratory of Biophysical Chemistry of the University of Groningen. This work was supported by The Netherlands Foundation of Chemical Research (CW) with financial aid from The Netherlands Foundation for Scientific Research (NWO).

RIJKSUNIVERSITEIT GRONINGEN

Molecular basis of two novel dehalogenating activities in bacteria

Structure/function relationships in an evolutionary context

Proefschrift

ter verkrijging van het doctoraat in de Wiskunde en Natuurwetenschappen aan de Rijksuniversiteit Groningen op gezag van de Rector Magnificus, dr. F. Zwarts, in het openbaar te verdedigen op vrijdag 30 januari 2004 om 14.15 uur

door

René Marcel de Jong

geboren op 22 maart 1976 te Heerenveen

Promotor: Prof. dr. B. W. Dijkstra

Beoordelingscommissie: Prof. dr. D. B. Janssen Prof. dr. A. E. Mark Prof. dr. J.B.F.N. Engberts

CONTENTS

CHAPTER 1: GENERAL INTRODUCTION ...... 9 EVOLUTION IN THE MICROBIAL WORLD...... 10 EVOLUTION AT THE MOLECULAR LEVEL...... 12 BACTERIAL DEHALOGENATING ACTIVITY TOWARDS ANTHROPOGENIC COMPOUNDS ...... 14 SCOPE OF THIS THESIS...... 17 REFERENCES ...... 20 CHAPTER 2: STRUCTURE AND MECHANISM OF BACTERIAL DEHALOGENASES: DIFFERENT WAYS TO CLEAVE A CARBON-HALOGEN BOND ...... 23 Abstract...... 23 INTRODUCTION ...... 24 DEHALOGENASE FAMILIES ...... 25 Haloalkane dehalogenases ...... 25 Haloacid dehalogenases ...... 27 4-Chlorobenzoyl-CoA dehalogenases...... 28 Haloalcohol dehalogenases ...... 29 3-Chloroacrylic acid dehalogenases...... 30 ON THE EVOLUTION OF DEHALOGENATING ACTIVITIES...... 33 REFERENCES ...... 35 CHAPTER 3: CRYSTALLIZATION AND PRELIMINARY X-RAY ANALYSIS OF AN ENANTIOSELECTIVE HALOHYDRIN DEHALOGENASE FROM AGROBACTERIUM RADIOBACTER AD1...... 39 Abstract...... 39 INTRODUCTION ...... 40 CRYSTALLIZATION, DATA COLLECTION AND PROCESSING ...... 42 REFERENCES ...... 44

CHAPTER 4: STRUCTURE AND MECHANISM OF A BACTERIAL HALOALCOHOL DEHALOGENASE: A NEW VARIATION OF THE SHORT- CHAIN DEHYDROGENASE/REDUCTASE FOLD WITHOUT AN NAD(P)H BINDING SITE ...... 47 Abstract ...... 47 INTRODUCTION ...... 48 RESULTS AND DISCUSSION ...... 50 Structure determination ...... 50 Overall structure of haloalcohol dehalogenase HheC...... 50 The halide binding site of haloalcohol dehalogenase HheC ...... 52 Product binding in haloalcohol dehalogenase HheC...... 54 Binding of a substrate mimic in haloalcohol dehalogenase HheC ...... 56 Structural relationships between haloalcohol dehalogenases and short-chain dehydrogenases/reductases ...... 56 Catalytic mechanism of haloalcohol dehalogenases...... 60 On the origin of haloalcohol dehalogenase activity...... 62 MATERIAL AND METHODS ...... 64 Crystallization of HheC...... 64 Structure determination and refinement ...... 65 Site-directed mutagenesis and kinetic characterization ...... 66 Structural alignment and sequence analysis...... 67 REFERENCES ...... 68 CHAPTER 5: STRUCTURAL BASIS FOR THE ENANTIOSELECTIVITY OF EPOXIDE RING OPENING REACTIONS CATALYZED BY HALOALCOHOL DEHALOGENASE HHEC...... 73 Abstract ...... 73 INTRODUCTION ...... 74 MATERIALS AND METHODS...... 77 Crystallization ...... 77 Data collection and refinement...... 79 Computational techniques...... 79 Free energy calculations ...... 80 RESULTS AND DISCUSSION ...... 81 Structure of an HheC:H2O:(R)-pNSO complex at 1.7 Å resolution ...... 81 Structure of an HheC:H2O:(S)-SO complex ...... 82 Structure of an HheC:H2O:(S)-pNSO complex at 1.9 Å resolution...... 85 Calculations of the relative Gibbs binding energy between (R)- and (S)-pNSO ...... 85 CONCLUSIONS...... 87 REFERENCES ...... 88 CHAPTER 6: THE X-RAY STRUCTURE OF TRANS-3-CHLOROACRYLIC ACID DEHALOGENASE REVEALS A NOVEL HYDRATION MECHANISM IN THE TAUTOMERASE SUPERFAMILY...... 91 Abstract ...... 91 INTRODUCTION ...... 92 METHODS ...... 94 Overall structure of CaaD...... 98 The active site of native CaaD ...... 98 The active site of inactivated CaaD...... 100

Mutagenesis of αArg-8 and αGlu-52 ...... 102 Similarity to other members of the 4-OT family ...... 103 Mechanism of inactivation of CaaD by 3-bromopropiolate...... 104 Conversion of trans-3-chloroacrylate by CaaD...... 106 The evolutionary origin of chloroacrylate dehalogenating activity ...... 107 REFERENCES ...... 108 CHAPTER 7: OUTLOOK ...... 111 INTRODUCTION ...... 112 WHAT ARE THE POSSIBLE HALIDE RELEASE PATHWAYS IN HHEC? ...... 112 CAN WE ENGINEER A REDOX ACTIVITY IN HALOALCOHOL DEHALOGENASE HHEC? ...... 114 CAN WE EXPERIMENTALLY EVOLVE DEHALOGENATING ACTIVITIES IN ENZYME SUPERFAMILIES? 115 REFERENCES ...... 117 CHAPTER 8: SUMMARY...... 119 INTRODUCTION ...... 120 DEHALOGENATION OF ANTHROPOGENIC HALOGENATED HYDROCARBONS BY BACTERIA...... 121 DIFFERENT WAYS TO CLEAVE A CARBON-HALOGEN BOND...... 121 MOLECULAR BASIS OF TWO NOVEL DEHALOGENATING ACTIVITIES IN BACTERIA...... 122 ON THE ORIGIN OF DEHALOGENATING ACTIVITIES IN BACTERIA...... 123 CHAPTER 9: NEDERLANDSE SAMENVATTING ...... 125 INLEIDING...... 126 DEHALOGENERING VAN ANTROPOGENE GEHALOGENEERDE KOOLWATERSTOFFEN DOOR BACTERIËN...... 127 VERSCHILLENDE MANIEREN OM EEN KOOLSTOF-HALOGEEN-BINDING TE VERBREKEN ...... 128 MOLECULAIRE GRONDSLAGEN VOOR TWEE NIEUWE DEHALOGENERINGSACTIVITEITEN ...... 129 DE OORSPRONG VAN DEHALOGENERINGSACTIVITEITEN IN BODEMBACTERIËN ...... 132 CHAPTER 10: NAWOORD...... 133

Chapter 1: General Introduction

All life evolves by the differential survival of replicating entities. - Richard Dawkins

There is grandeur in this view of life, with its several powers, having been originally breathed into a few forms or into one; and that, whilst this planet has gone cycling on according to the fixed law of gravity, from so simple a beginning endless forms most beautiful and most wonderful have been, and are being, evolved. - Charles Darwin

Evolution on an ancient earth is as well established as our planet's shape and position. Our continuing struggle to understand how evolution happens - the "theory of evolution" - does not cast our documentation of its occurrence - the "fact of evolution" - into doubt. - Stephen Jay Gould

Introduction It is generally believed that evolution is at the heart of species diversification of living organisms, a hypothesis that was first stated by Charles Darwin in his revolutionary work “The Origin of Species”. Numerous descriptions of different animals, plants, and microbes have shown their phenotypic similarities, manifested, for example, by the similarities in organs, leaves, and flowers, and on the cellular level in their cell organelles. The ongoing revelation of the secrets of the genetic code, which started 50 years ago with the publication of the structure of DNA (Figure 1) (Watson and Crick, 1953), ultimately showed that evolutionary relationships between species are even more evident on the genetic level, even when species have very dissimilar phenotypes (Kirkness et al., 2003). Central to the current view of Darwinian evolution and species differentiation are: (i) mutation/recombination of the genetic code and the corresponding phenotypic changes (mutation); (ii) differential survival of altered individuals in the population (selection); and (iii) the propagation of beneficial changes in the population by (a)sexual replication (amplification). Despite this concise theoretical concept, the mechanisms that gave rise to the current species diversity are poorly understood. This partly lies in the tremendously large timescale of evolutionary changes to occur in higher organisms, making it impossible to get the slightest idea of the intermediates along the pathway. Often, fossils are the only proof of the past existence of such 'missing links' that could support the hypothesized evolutionary trajectories between different species. For example, fossils were of great value to substantiate the hypothesis of the evolution of birds from reptiles (Xu et al., 2003), and also have revealed important clues to human evolution (White et al., 2003).

Evolution in the microbial world

The theory of evolution has long been in the realm of theoretical biology, suffering from the inaccessibility to empirical experiments to support hypotheses. Nowadays, sequenced genomes from various species from all four kingdoms of life provide a huge amount of data to study evolutionary relationships on the nucleotide and α-amino acid sequence level. Moreover, molecular biology techniques have given access to new experimental approaches using microbial or viral species. Examples of the evolutionary adaptations in microbes are present all around us, and provide useful model systems. Striking examples are the emergence of antibiotic resistance in various pathogenic bacteria (McDermott et al., 2003), the adaptation of the HIV virus to anti-viral drugs (Hanna, 2002), and very recently the adaptation of an animal virus to infect humans,

10 leading to Severe Acute Respiratory Syndrome (SARS) (Guan et al., 2003). Bacteria are also useful to monitor natural adaptation in the laboratory, as they divide on average every 15 minutes. Consequently, experimental evolution of 1000 generations of a cultured bacterium requires only ten days (Riley et al., 2001). Such empiric approaches have not often been reported, but this may change in view of recently communicated experiments with very stimulating results (Rainey and Rainey, 2003; Velicer and Yu, 2003).

Figure 1. A surface representation of two turns of double-stranded DNA (top). The transparent surface shows the schematic structure of the chain of inward-directed bases that form the genetic code. The lower part illustrates how base changes in a DNA strand may lead to differences in the transcribed mRNA product. Depending on the changes in the 3-letter codons of the mRNA, this may lead to the incorporation of different amino acids into the translated protein, which subsequently may result in functional differences in the folded protein or enzyme.

The ability of microbial species to rapidly adapt to a changing environment, their high evolvability (Conrad, 1990; Kirschner and Gerhard, 1998), is also apparent from the microbial diversity found in nature. Microbial species represent the largest species diversity on earth. Apart from soil and surface water, microbes inhabit almost every noxious environment on earth, from the boiling water of hot springs to the Antarctic sea ice (Rothchild and Mancinelli, 2001; Thomas and Dieckmann, 2002). Microbial communities may contain a tremendous diversity of species, and their composition is of complex and dynamic nature (Torsvik and Øvreas, 2002). One gram of soil can harbour up to 10,000 prokaryotic species, of which up to 99% are still undefined (Torsvik et al., 1998). The community metagenome (Handelsman et al., 1998) may contain over 1012 different genes, which is in large contrast to only 104 different proteins and enzymes that have currently been described in the literature (Seffernick and Wackett, 2001). The microbial world thus not only represents the largest species diversity, but millions of years of evolution also resulted in by far the largest sequence diversity on earth. Microbial species thus provide an infinite source of gene sequences, encoding functional components of numerous catabolic pathways and regulation systems, of which many remain to be discovered.

Evolution at the molecular level

Despite our very narrow view on the actual extent of the diversity of gene sequences and the translated amino acid sequences in nature, it is clear that the number of different protein folds is limited (Chothia et al., 2003). Structure determination of various proteins showed that in contrast to an infinite sequence space, fold space is estimated to contain between 650 and 10,000 different folds (Figure 2) (Chothia et al., 2003). The folded states that specific amino acid sequences adopt in a cell are critical to understand the molecular mechanism of its function. Consequently, mutations in a gene that result in an altered amino acid sequence may directly affect its function through the structural perturbation of its folded state. This direct link between gene/protein sequence and protein structure/function thus suggests a mechanism of evolution on the molecular level, in which mutation of a gene may lead to functional drift of the translated protein. This may have resulted, for example, in the various superfamilies of enzymes that exist, of which the similarly folded members have distinctly different catalytic activities (Murzin, 1996; Babbitt and Gerlt, 1997). Often, the different representatives share little sequence identity, but the similar fold and partly conserved catalytic residues suggest distant evolutionary

12 relationships. The adaptive routes along which the different activities emerged in a superfamily are, however, poorly understood (see e.g. Gerlt and Raushel, 2003). These difficulties are partly a consequence of the absence of 'protein fossils'.

Figure 2. Schematic representations of a few different protein folds that have been determined by X- ray crystallography. A) The trimeric structure of the integral outer membrane protein TolC, a component of the drug efflux pump of Escherichia coli, consisting of a membrane-embedded β-barrel and an α-helical barrel domain (Koronakis et al., 2000). B) The tetrameric structure of the β- galactosidase lacZ, a sugar-processing enzyme of E. coli, consisting of four identical subunits of mostly β-strands (Juers et al., 1994). C) The dimeric structure of the λ repressor-operator complex, an α-helical regulating protein that is bound to double-stranded DNA. The specific DNA sequence is recognized by the α-helices that bind in the major grooves of the DNA, which regulates the transcription of a neighboring gene (Beamer and Pabo, 1992). D) The tetrameric structure of haloalcohol dehalogenase HheC from Agrobacterium radiobacter AD1, a dehalogenating enzyme that plays an important role in degradation of toxic haloalcohols in this bacterium (de Jong et al., 2003). E) The monomeric structure of haloalkane dehalogenase DhlA from Xanthobacter autothrophicus GJ10, which plays a role in the degradation of toxic haloalkanes (Franken et al., 1991).

Bacterial dehalogenating activity towards anthropogenic compounds

From an evolutionary point of view, enzymes that facilitate the growth of certain bacteria on anthropogenic halogenated compounds are especially intriguing, as some of them may have only recently evolved by adaptation of related enzymes in response to elevated concentrations of these compounds in the environment. Although many halogenated compounds occur naturally (Oberg, 2002), compounds like 1,2- dibromoethane and 1,3-dichloro-2-propanol are presumed to be xenobiotic, meaning that they were absent in the environment before their industrial production (Figure 3). The (geno)toxic properties of some man-made halogenated compounds may have put a large evolutionary pressure on bacteria to evolve enzymes that facilitate cleavage of the carbon- halogen bond, and release the halogen as a harmless halide ion. Various catabolic pathways that depend on such dehalogenases have been identified in bacteria isolated from contaminated soil or water samples, in which they convert specific halogenated compounds into alternative growth substrates for the bacterium. The clearest example known to date of a recently evolved bacterial enzyme that converts a xenobiotic halogenated substrate is atrazine chlorohydrolase, which catalyzes the hydrolytic cleavage of a carbon-halogen bond in atrazine, a herbicide of which over 2 billion pounds have been applied to soils globally (Seffernick and Wackett, 2001). The amino acid sequence of atrazine chlorohydrolase is 98% identical to the enzyme melamine deamidase, a member of a large aminohydrolase enzyme superfamily. The high identity of the two enzymes, and the knowledge that melamine deamidase also converts a xenobiotic substrate (Seffernick and Wackett, 2001), evidence a recent evolutionary link between the two. Unfortunately, structural information on the dehalogenase that could reveal a structural basis for its activity is not available. Characterization of several other dehalogenases from bacteria revealed a wide variety of different chemical strategies that are used in nature to facilitate cleavage of carbon-halogen bonds (Figure 4) (Fetzner, 1998; Janssen et al., 2001; van Pee dan Unversucht, 2003). Several anaerobic halorespiring bacteria contain reductive dehalogenases that enable them to use tetrachloroethene, rather than oxygen, as the final electron acceptor in the respiratory chain (Chen, 2003). Aerobic bacteria, instead, may contain mono- or dioxygenases to oxidatively dehalogenate small haloalkenes, haloalkanes and some aromatic compounds (Fetzner, 1998). A third dehalogenation

Figure 3. A small selection of known halogenated compounds, their molecular structure, and their respective anthropogenic or natural origins.

Figure 4. A selection of different bacterial dehalogenases, and the dehalogenation reactions that they facilitate.

strategy is represented by dichloromethane dehalogenase, which uses a gluthathione cofactor to dehalogenate dichloromethane via a thiolytic substitution mechanism (Leisinger et al., 1994). Most of the characterized bacterial dehalogenases, however, utilize cofactor- independent substitution mechanisms that proceed via a covalently bound intermediate with a nucleophilic aspartate, which is subsequently hydrolyzed by a water molecule. In contrast to the reductive and thiolytic dehalogenases, a wealth of structural information is available for several of those hydrolytic dehalogenases, which revealed that they belong to unrelated enzyme superfamilies. However, no close sequence homologues are presently known that could identify the likely precursor enzymes of the different dehalogenases. Apart from the outlined evolutionary point of view, dehalogenases are also intriguing from a (bio)chemical perspective, as they facilitate chemical reactions that normally are often very unfavorable to occur spontaneously in aqueous media. It was long believed that various industrially-produced halogenated hydrocarbons were 'undegradable', but over the past five decades it became clear that even the most notorious and persistent ones can be enzymatically degraded by some bacteria (Bunge et al., 2003). Isolation and culturing of such bacteria offered exciting opportunities for bioremediation, as they could provide potential 'green' solutions to inexpensively restore contaminated environments, or to clean industrial waste-water streams of the chemical industry (Swanson, 1999). Biochemical and structural information on the isolated dehalogenases can provide important insights into the molecular basis of enzymatic dehalogenation reactions, and provide the opportunity to improve or alter their catalytic properties by structure-based design.

Scope of this thesis

This thesis describes the X-ray crystallographic elucidation of the molecular structures of two novel unrelated bacterial dehalogenases, the haloalcohol dehalogenase HheC from Agrobacterium radiobacter AD1, and a trans-3-chloroacrylic acid dehalogenase (CaaD) from Pseudomonas pavonaceae 170. Structures of the native enzymes, and of complexes of the enzymes with products, substrate analogues, or inhibitors provide conclusive insights into the mechanistic details that underlie their catalytic activities. Furthermore, structural comparison with structurally related enzymes with different catalytic activities reveals new insights into their possible evolutionary origins.

Chapter 2 provides an extensive structural comparison of the structures and the novel dehalogenation strategies of HheC and CaaD, which fundamentally differ from the structures and mechanisms of previously characterized haloalkane -, haloacid -, and 4- chlorobenzoyl-Coenzyme A dehalogenases. The structures and mechanisms of these dehalogenases are further compared with their closest structural relatives. Whereas the evolutionary origin of most of the other dehalogenases remains largely unclear, the structural information on haloalcohol dehalogenase and trans-3-chloroacrylate dehalogenase that have become available through these studies offer convincing evidence for their likely origins. Chapter 3 reports the crystallization procedures of haloalcohol dehalogenase HheC, which plays a role in the degradation of haloalcohols in A. radiobacter AD1. HheC catalyzes the dehalogenation of vicinal haloalcohols like 2,3-dichloro-1-propanol via an one-step intramolecular substitution mechanism, yielding the corresponding epoxide and HCl. In Chapter 4, X-ray structures of haloalcohol dehalogenase HheC are presented, which reveal the molecular basis of the catalyzed dehalogenation reaction. The results further show that HheC is structurally and mechanistically related to a widespread family of NAD(P)H-dependent redox enzymes, the short-chain dehydrogenases/reductases (SDR family). X-ray structures of HheC with bound products and a substrate analogue showed that catalysis depends on a conserved Ser-Tyr-Arg catalytic triad. The SDR NAD(P)H- binding site has changed, however, into a halide-binding site that supports the cofactor- independent dehalogenation reaction. Structure-based sequence alignments further evidenced that the six haloalcohol dehalogenases that are currently known have evolved from two different NAD-binding SDR enzymes, rather than from NAD(P)-binding enzymes. There is considerable interest in haloalcohol dehalogenases from a biocatalytic point of view, as they may have promising applications in the green synthesis of valuable enantiopure building blocks of biologically active compounds (de Vries and Janssen, 2003). Haloalcohol dehalogenase HheC catalyses the highly enantioselective dehalogenation of vicinal haloalcohols to epoxides (50 < E < 150), as well as the reverse reaction, the enantioselective and β-regioselective nucleophilic ring opening of epoxides by halides, but also pseudo-halides like azide and cyanide (E > 200). In chapter 5, the X- ray structures of complexes of HheC with the unfavored S-enantiomers of styrene oxide and p-nitro-styrene oxide are presented. Comparison with the X-ray structure of HheC in complex with the catalytically favoured R-enantiomer of styrene oxide (R-SO) and a

18 chloride ion, reveals that the unfavored S-enantiomers of epoxides can also bind in the active site, but in a non-productive ‘upside-down’ manner. Comparison of the different complexes and calculations of the Gibbs energy difference between the productive and non-productive states provide a structural explanation for the higher enantioselectivity of epoxide ring opening reactions with respect to the dehalogenation reactions catalyzed by HheC. In chapter 6, I present X-ray structures of the trans-3-chloroacrylic acid dehalogenase CaaD from P. pavonaceae 170, of the native enzyme and a form that is covalently inactivated by a mechanism-based inhibitor. The results provide conclusive evidence that CaaD cleaves an sp2-hybridized carbon-halogen bond via a Michael addition of water to trans-3-chloroacrylate, a degradation product of the soil fumigant 1,3- dichloropropene. Subsequent decomposition of the product, an unstable halohydrin, results in the release of malonate semialdehyde and HCl. CaaD is related to the well- studied Pseudomonas enzyme 4-oxalocrotonate tautomerase (4-OT). Whereas this tautomerase uses an N-terminal catalytic proline (Pro1) as a base to transfer protons in tautomerization reactions, the Pro1 in the dehalogenase is acidic, stabilized in a hydrophilic active site. Together with a buried glutamate base, the acidic proline facilitates a hydratase activity, rather than a tautomerase activity. Intriguingly, 4-OTs have a rudimentary dehalogenation activity, suggesting that CaaD may have evolved from a 4-OT precursor. Finally, chapter 7 provides an outlook on future plans to use the available structural information on haloalcohol dehalogenase HheC to design an NAD-dependent redox activity in the enzyme, thus restoring the activity of its precursor family. Other plans aim to resolve remaining questions of the pathway(s) that are used by halides and nucleophilic anions to exit or enter the halide-binding site.

References

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Chapter 2: Structure and mechanism of bacterial dehalogenases: Different ways to cleave a carbon-halogen bond

René M. de Jong & Bauke W. Dijkstra

Abstract

Dehalogenases make use of fundamentally different strategies to cleave carbon-halogen bonds. The structurally characterized haloalkane dehalogenases, haloacid dehalogenases, and 4- chlorobenzoate-CoA dehalogenases use substitution mechanisms that proceed via a covalent aspartyl intermediate. Recent X-ray crystallographic analysis of a haloalcohol dehalogenase and a trans-3-chloroacrylic acid dehalogenase provide detailed insight into a different intramolecular substitution mechanism and a hydratase-like mechanism, respectively. The available information on the various dehalogenases supports different views on the possible evolutionary origins of their activities.

Published in: Current Opinion in Structural Biology 2003, 13, 722-730.

Introduction

From the beginning of the past century, halogenated hydrocarbons have been extensively applied in industry and agriculture. Decades after the start of their widespread use, evidence started to accumulate that some of these xenobiotic halogenated compounds are highly toxic. Notorious examples are dioxins and polychlorinated biphenyls (PCBs), but also halogenated solvents like for instance 1,2-dichloroethane have been classified as probably carcinogenic to humans (Safe, 1994; Salovsky et al., 2002). Because of these noxious properties, many of these organohalogens have now been banned from use and have been replaced by environmentally less harmful compounds. Halogenated compounds are, however, not only of anthropogenic origin. Over 1500 organohalogens are known that are produced naturally (Odberg, 2002; Balschmitter, 2003). They range from volatile compounds such as methylchloride to antibiotics like vancomycin and chloramphenicol. Insight into the enzymes that incorporate halogens into organic compounds is rather limited (van Pée and Unversucht, 2003). This strongly contrasts with the large amount of detailed information on enzymes that degrade halogenated hydrocarbons, releasing the halogens as halide ions. In this review we focus on bacterial dehalogenases. These enzymes make use of a variety of distinctly different catalytic mechanisms to cleave carbon-halogen bonds. X-ray structures of haloalkane dehalogenases, haloacid dehalogenases, and 4-chlorobenzoyl- CoA dehalogenase demonstrated the power of substitution mechanisms that proceed via a covalent aspartyl intermediate (Verschueren et al., 1993; Benning et al., 1996; Hisano et al., 1996; Ridder et al., 1999). Recent structural characterizations of a haloalcohol dehalogenase and a trans-3-chloroacrylic acid dehalogenase reveal the details of two other elegant catalytic strategies, which exploit catalytic mechanisms from homologous enzymes in a different chemical context (de Jong et al., 2003; de Jong et al., submitted). The available information on the various dehalogenases supports different views on the possible origins of their individual activities towards presumed anthropogenic halogenated substrates.

Figure 1. The bacterial degradation routes of four different halogenated compounds, in which the five families of dehalogenases discussed in this chapter facilitate the enzymatic cleavage of the carbon- halogen bonds.

Dehalogenase families

Haloalkane dehalogenases

Haloalkane dehalogenases cleave the carbon-halogen bond in halogenated aliphatic hydrocarbons (Figure 1A). The haloalkane dehalogenase from Xanthobacter autotrophicus GJ10 was the first dehalogenase of which the crystal structure was determined (Franken et al., 1991), but later also other structures have become available (Newman et al., 1999; Oakley et al., 2002). The structure consists of two domains, a main domain and a cap domain (Figure 2A). The main domain consists of a mostly parallel eight-stranded β-sheet (only the second β-strand is antiparallel), connected by α-helices on both sides of the β-sheet. The cap domain is α-helical. This fold is the hallmark of the superfamily of α/β-hydrolases, to which, besides the haloalkane dehalogenases, also lipases, esterases, carboxypeptidases, and acetylcholinesterases belong (Ollis et al., 1992; Heikinheimo et al., 1999; Nardini & Dijkstra, 1999). These latter enzymes catalyze the hydrolysis of ester and amide bonds via a two-step nucleophilic acyl substitution mechanism similar to that of serine proteases (see e.g. Dodson and Wlodawer, 1998;

Jaeger et al., 1999). In the first step, the serine or cysteine of a Ser/Cys-His-Asp/Glu catalytic triad functions as a nucleophile that attacks the sp2-hybridized carbonyl carbon atom of the scissile ester/amide bond, resulting in a covalently bound ester intermediate. Subsequently, a water molecule activated by the His/Asp or His/Glu pair hydrolyzes this ester intermediate in the second step. Haloalkane dehalogenases use a very similar two-step catalytic mechanism, except that the nucleophile is an aspartate instead of a serine/cysteine residue, which, in 3 an SN2 type substitution mechanism, attacks the halogen-bearing sp -hybridized carbon atom. This leads also to a covalently bound ester intermediate that can be easily hydrolyzed by attack of a His-activated water molecule on the Cγ atom of the aspartate (Figure 2C) (Verschueren et al., 1993). Thus, the substitution of the serine/cysteine of an α/β-hydrolase by an aspartate confers on the enzyme an essential prerequisite to hydrolyze carbon-halogen bonds in haloalkanes. In addition, haloalkane dehalogenases contain a halide-binding site to facilitate the dehalogenation reaction.

Figure 2. A) Structure of haloalkane dehalogenase from X. autotrophicus (Franken et al., 1991), showing the nucleophilic aspartate and a bound chloride ion. B) Dimeric structure of haloacid dehalogenase with a covalently bound intermediate (Ridder et al., 1999). C) Schematic of the catalytic mechanisms of haloalkane and haloacid dehalogenases.

During recent years, computational studies have unravelled the details of the catalytic mechanism. In the enzyme, the nucleophilic aspartate is nearly entirely positioned in a “near attack conformation”, with the attacking oxygen of the aspartate in line with the C-Cl bond of the substrate, at a distance favorable for attack of the halogen-bearing carbon atom. The correct positioning accounts, however, only for 25% to the lowering of the transition state free energy (Shurki et al., 2002; Hur et al., 2003). The remaining 75% probably originates from contributions of the halide-binding residues to transition state stabilization (Boháč et al., 2002) and from the half-hydrophilic/half-hydrophobic nature of the active site, which facilitates both the activation of the nucleophile and the departure of the leaving group. Computational quantification of these contributions is presently beyond reach (Hur et al., 2003).

Haloacid dehalogenases

Haloacid dehalogenases catalyze the hydrolysis of α-halogenated carboxylic acids, such as 2-chloroacetate, which is an intermediate in the degradation of 1,2- dichloroethane (Figure 1A). They are members of the haloacid dehalogenase (HAD) superfamily (Koonin and Tatusov, 1994; Ridder et al., 1999), to which also magnesium- dependent phosphatases and P-type ATPases belong. The haloacid dehalogenases are dimers with two- or three-domains per subunit: a core domain with a Rossmann-fold-like six-stranded parallel β-sheet flanked by five α-helices, a sub-domain consisting of a four- helix bundle, and, in some enzymes, a dimerization domain of two anti-parallel α-helices (Figure 2B) (Hisano et al., 1996; Ridder et al., 1997). This fold is completely different from the α/β-hydrolase fold of the haloalkane dehalogenases. Like the latter enzymes, the haloacid dehalogenase utilizes an aspartate-based catalytic mechanism that proceeds via a covalent intermediate (Figure 2C), but there is no histidine to activate a nucleophilic water molecule. Also, the halide-binding site is very different (Ridder et al., 1999). How the water molecule is activated is not known, but it has been suggested that another aspartate in the active site fulfils this function (Ridder et al., 1999). Intriguingly, at least one other unrelated haloacid dehalogenase family exists (Hill et al., 1999; Marchesi & Weightman, 2003), which was shown to bypass the classic covalent ester intermediate (Nardi-Dei et al., 1999). This suggests that enzymes with different haloacid dehalogenating strategies have evolved from different enzyme precursors, but unfortunately, crystallographic data that could provide structural evidence on the mode of action of these type II haloacid dehalogenases is still unavailable.

4-Chlorobenzoyl-CoA dehalogenases

The 4-chlorobenzoyl-Coenzyme A (CoA) dehalogenase from Pseudomonas sp. strain CBS-3 is a homotrimer, of which each subunit folds into two domains. The N- terminal domain contains a 10-stranded β-sheet, forming two nearly perpendicular layers, which are flanked by α-helices (Figure 3A). The C-terminal domain is composed of three amphiphilic α-helices, and is primarily involved in trimerization. Thorough characterization of the enzyme has revealed the details of the mechanism by which a halogen is displaced from the aromatic ring of 4-chlorobenzoate, a degradation product of PCBs (Figure 1D) (Benning et al., 1996; Luo et al., 2001; Dong et al., 2002). The mechanism also proceeds via a covalent aspartyl intermediate. First, the substrate is ligated to CoA by 4- chlorobenzoate CoA ligase. The CoA-ligated product is then bound by the 4-chlorobenzyl- CoA dehalogenase, with the enolate anion of the thioester link of the CoA-ligated substrate stabilized by two backbone NH groups. This induces a partially positive charge on the halogen-bearing carbon atom, which makes it susceptible to nucleophilic attack by the aspartate (Figure 3B) (Luo et al., 2001). The SNAr-type substitution results in a covalent Meisenheimer intermediate, which was recently confirmed by Raman spectroscopic measurements (Dong et al., 2002). Chloride is expelled upon restoration of the aromatic ring system, producing a second arylated enzyme intermediate that is subsequently hydrolyzed by attack on the Cγ atom of the nucleophilic aspartate by a histidine-activated water molecule, yielding 4-hydroxybenzoyl-CoA (Zhang et al., 2001; Lau et al., 2002). The stabilization of the acyl-CoA enolate anion intermediate is shared with other hydratases, isomerases and thioesterases of the crotonase (or enoyl-CoA hydratase) superfamily (Holden et al., 2001) but the nucleophilic aspartate is only present in the dehalogenase. Like in haloalkane and haloacid dehalogenases, the aspartate confers to the enzyme its unique capability to hydrolyze the bond between a halogen and an aromatic carbon atom.

Figure 3. A) Trimeric structure of 4-chlorobenzoyl-coenzyme A dehalogenase (Benning et al., 1996), showing the nucleophilic aspartate, a bound calcium ion, and the product 4-hydroxybenzoyl- coenzyme A. B) Schematic of the catalytic mechanism of 4-chlorobenzoyl-coenzyme A dehalogenase.

The enzymes discussed above illustrate the power of aspartate-mediated substitution mechanisms to hydrolyze organohalogens. However, several other dehalogenases exist that use completely different catalytic strategies. For instance, halorespiring anaerobic bacteria contain reductive dehalogenases, but unfortunately their structures are not yet known (Wohlfarth and Diekert, 1997; Furukawa, 2003). In contrast, X-ray structures of a haloalcohol dehalogenase and the trans-3-chloroacrylic acid dehalogenase CaaD have recently provided detailed insight into two other fundamentally different dehalogenation strategies.

Haloalcohol dehalogenases

Several bacteria contain enzymes that are able to displace a halogen from a vicinal haloalcohol substrate. One such enzyme is the haloalcohol dehalogenase HheC from Agrobacterium radiobacter AD1, which plays a role in the degradation of 1,3-dichloro- 2-propanol (Figure 1B). HheC is a homotetrameric enzyme (Figure 4A): the monomer is homologous to that of members of the widespread short-chain dehydrogenase/reductase (SDR) family. SDR-enzymes are redox enzymes that catalyze alcohol-ketone conversions

29 of a wide variety of alcohols, steroids and sugars (Filling et al., 2002; Oppermann et al., 2003). They have the well-known dinucleotide-binding Rossmann fold (Rossmann et al., 1974) and a Ser-Tyr-Lys/Arg catalytic triad. HheC shares the fold and catalytic triad of the SDR-family, but lacks the characteristic dinucleotide-binding Gly-X-X-X-Gly-X-Gly motif (Figure 4D) (van Hylckama Vlieg et al., 2001; de Jong et al., 2003). Instead, several larger residues replace the smaller ones of the motif, thus filling up the space of the cofactor- binding site. In this way, a spacious halide-binding site is created (Figure 4B). The enzyme uses an intramolecular SN2-type substitution mechanism, in which the secondary hydroxyl group of the haloalcohol substrate is deprotonated by the tyrosine of the conserved Ser- Tyr-Arg catalytic triad acting as a base (Figure 4C). The hydroxyl oxygen concomitantly substitutes the vicinal halogen to yield the corresponding epoxide product, a proton and a chloride ion. Thus, while the aspartate-dependent dehalogenases discussed above catalyze a two-step hydrolysis of the carbon-halogen bond, the presence of a hydroxyl function vicinal to the halogen allows the one-step, intramolecular substitution of the halogen by the hydroxyl.

3-Chloroacrylic acid dehalogenases

Like 4-chlorobenzoyl-CoA dehalogenases, the chloroacrylic acid dehalogenases displace a halogen from an sp2-hybridized carbon atom. The cis- and trans-3-chloroacrylic acid dehalogenases from Pseudomonas pavonaceae 170 (Poelarends et al., 2001) play a role in the degradation of 1,3-dichloropropene, a compound applied in agriculture to kill plant-infecting nematodes (Figure 1C). After conversion of trans-1,3-dichloropropene by a haloalkane dehalogenase, oxidation of the trans-1-chloro-3-hydroxypropene product yields trans-3-chloroacrylate. The trans-3-chloroacrylic acid dehalogenase (CaaD) converts this product into malonate semialdehyde, a chloride ion and a proton. Subsequent decarboxylation of malonate semialdehyde yields acetaldehyde, which can be used as an alternative growth substrate. The X-ray structures of the native and an inactivated form of trans-3-chloroacrylic acid dehalogenase (CaaD) have recently been solved (de Jong et al., submitted). The enzyme is a trimer of αβ heterodimers (Figures 5A, 5B). Its fold is similar to that of 4- oxalocrotonate tautomerase (4-OT), a member of the tautomerase superfamily, which contains isomerases and tautomerases (Whitman, 2003). However, whereas in the isomerases and tautomerases the catalytic Pro-1 is in a hydrophobic environment, in CaaD the corresponding Pro-1β is in a hydrophilic active site near a buried glutamate. This

30 has a major effect on the pKa of the proline residue: in 4-OT the proline has a strongly reduced pKa (Czerwinski et al., 2001), whereas in CaaD the pKa is normal. This has consequences for the catalytic role of the proline: in the isomerases and tautomerases it functions as a general base, but in CaaD it functions as a general acid.

Figure 4. A) Tetrameric structure of haloalcohol dehalogenase (de Jong et al., 2003) with a bound chloride ion and a bound epoxide product. B) Detailed view of the halide binding site. Trp249* comes from another subunit. Dashed lines indicate hydrogen bonds. C) Schematic of the catalytic mechanism of haloalcohol dehalogenase. D) Sequence comparison of the haloalcohol dehalogenases HheC, HheA, and HheB with 4 SDR-family members indicated by their PDB entry number. 1FMC is an NAD-dependent enzyme, while 1YBV, 1AE1, and 2AE2 are NADP-dependent. An aspartate or glutamate in one of the boxed positions is indicative of an NAD-dependent enzyme, whereas NADP- dependent enzymes have 1 or 2 arginines or lysines in different positions. NAD-dependent SDR subfamilies are defined among others by the position of the aspartate (position 1, 2 or 3 in the box).

Figure 5. A) Heterohexameric structure of trans-3-chloroacrylic acid dehalogenase with a covalently bound malonyl adduct to Pro-1β (de Jong et al., in press). B) Structure of the trans-3-chloroacrylic acid dehalogenase monomer with a covalently bound malonyl adduct to Pro-1β. C) Stereo view of the active site showing the bound adduct. The red and pink main chains indicate chains from the α and β subunits, respectively. D) Schematic of the catalytic mechanism of trans-3-chloroacrylic acid dehalogenase.

The role of the Pro-1β in CaaD became clear from the crystal structure of CaaD inactivated by the suicide substrate 3-bromopropynoate (Figure 5C) (de Jong et al., submitted). This compound inactivates the enzyme by forming a covalent malonyl adduct with the proline residue. This modification is the result of the hydration of the carbon- carbon triple bond of the inhibitor via a Michael addition, catalyzed by the active site glutamate, which activates a water molecule, and by the proline, which donates a proton. The reactivity of the triple bond is increased by electrostatic interactions of its conjugated carboxylate group with two arginine residues. In the case of 3-bromopropynoate the product rearranges to a reactive acyl bromide, which forms a covalent bond with the deprotonated proline. In contrast, the substrate trans-3-chloroacrylate is converted to an instable halohydrin that decomposes to form malonate semialdehyde, a chloride ion and a proton (Figure 5D). Thus, the conjugated system of the carbon-carbon double bond and the carboxylate group enables the dehalogenase to hydrate the substrate via a conjugate addition reaction, followed by spontaneous dehalogenation of the product. The active site glutamate and proline thus form the basis of a previously unidentified hydratase activity in the tautomerase superfamily.

On the evolution of dehalogenating activities

The structural characterization of several bacterial dehalogenases has provided detailed insight into their evolutionary relationship to other enzyme families. Some dehalogenases may have evolved from superfamily members with distinctly different activities, analogous to the creation of low-level crotonase activity in 4-chlorobenzoyl-CoA dehalogenase by a double glutamate mutation (Xiang et al., 1999). Alternatively, dehalogenation activity may emerge alongside an original enzyme activity. This is illustrated by the bacterial muconate lactonizing enzymes, some of which can also dehalogenate chlorinated muconates. A few amino acid substitutions can account for this dehalogenating activity (Kajander et al., 2003). Although these case studies nicely illustrate that minimal changes can drastically alter enzyme activities, they do not provide compelling evidence on the origins of the various isolated dehalogenases. The recent adaptation of a precursor enzyme in response to large concentrations of a xenobiotic halogenated substrate may seem a reasonable mechanism for the emergence of these activities, but seems difficult to reconcile with the worldwide spread of some of the enzymes. Conserved gene clusters containing haloalkane-utilizing catabolic

33 pathways have been isolated from bacterial strains from three different continents (Poelarends et al., 2000). Although horizontal gene transfer and/or integrase-dependent gene acquisition can help in the local spread in bacteria (Poelarends et al., 2000), a global diffusion by this mechanism seems unlikely. However, the presence of haloalkane dehalogenases in various parasitic Mycobacterium strains, which colonize both animal tissues and the free environment, could suggest a possible role of parasitic microorganisms in a worldwide distribution mechanism (Jesenská et al., 2000). The global distribution of these enzymes could also support a pre-industrial origin. Many different organohalogens occur naturally, suggesting that corresponding dehalogenating activities exist in nature. Even in the case of highly toxic dioxins an anaerobic bacterium has been isolated that can grow on them (Bunge et al., 2003). Dioxins were long believed to be exclusively produced by the recent industrial activity of mankind (Alcock and Jones, 1996), but they have entered the environment also by natural processes (Hoekstra et al., 1999) and by the pre-industrial domestic burning of peat (Meharg and Killham, 2003). Although all dehalogenases that have been structurally characterized up to now are members of existing enzyme superfamilies, their sequences are highly divergent from superfamily members without dehalogenase activity. However, a structure-based sequence alignment of haloalcohol dehalogenases and SDR enzymes demonstrates a significant sequence identity of up to 30 to 35 % (de Jong et al., 2003). Two families of haloalcohol dehalogenases can be discerned that are related to different coenzyme-based SDR subfamilies, indicating that the two dehalogenase families independently originated from two different NAD-binding precursors, rather than NAD(P)H-binding precursors (Figure 4D). It is conceivable, that the haloalcohol could have bound in the active site of such an SDR enzyme, whereupon the catalytic tyrosine fortuitously facilitated the intramolecular substitution of the halogen by deprotonating the intramolecular hydroxyl group. The dehalogenase activity could have been improved as a result of evolutionary optimization of this rudimentary promiscuous activity. Catalytic promiscuity has been discovered in various other enzyme superfamilies that use similar structural features in a different chemical context (O'Brien and Herschlag, 1999). Strikingly, the closest relatives of 3-chloroacrylic acid dehalogenase, the tautomerase 4-OT and its homologue YwhB, have a promiscuous dehalogenating activity towards trans-3-chloroacrylic acid (Whitman, 2003). It would be interesting to see whether some existing members of the SDR family display a low haloalcohol dehalogenation activity.

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Chapter 3: Crystallization and preliminary X-ray analysis of an enantioselective halohydrin dehalogenase from Agrobacterium radiobacter AD1

René M. de Jong, Henriëtte J. Rozeboom, Kor H. Kalk, Lixia Tang, Dick B. Janssen & Bauke W. Dijkstra

Abstract

Halohydrin dehalogenases are key enzymes in the bacterial degradation of vicinal halopropanols and structurally related nematocides. Crystals of the enantioselective halohydrin dehalogenase HheC from Agrobacterium radiobacter AD1 have been obtained at room temperature from hanging drop vapor diffusion experiments against 50 - 70% saturated ammonium-sulfate solution at pH 6.5 to 7.3. The crystals belong to space group P41212 or P43212 with cell axes a = b = 104.5 Å and c = 121.4 Å, and contain two monomers in the asymmetric unit. The crystals diffract to 3.0 Å resolution with X-rays from a Cu Kα rotating anode generator.

Published in: Acta Crystallographica 2002, D58, 176-178.

Introduction

Haloorganic compounds are widespread in nature. While most of these compounds originate from both biotic and abiotic natural sources, many short-chain halogenated compounds are produced industrially on a large scale. The intentional and accidental introduction of several of these latter compounds into the environment has caused important pollution problems, because of their persistency and toxicity. Fortunately, bacteria have been found that are able to degrade some of these halogenated hydrocarbons and even use them as a growth substrate. Often, these bacteria use dehalogenases to break the bond between a carbon and a halogen. It is no surprise that some of these enzymes have been extensively studied in view of their detoxifying properties (Fetzner and Lingens, 1994; Fetzner, 1998). Enzymes that convert vicinal halohydrins form a special dehalogenase subclass. These halohydrin dehalogenases are also referred to in the literature as haloalcohol dehalogenases, halohydrin hydrogen-halide lyases or halohydrin epoxidases. They catalyze the reversible conversion of a vicinal haloalcohol such as 2,3-dichloro-1-propanol to its corresponding epoxide by the intramolecular nucleophilic substitution of the halogen by the vicinal hydroxyl group, thereby releasing a proton and a halide ion (Figure 1) (Castro and Bartnicki, 1968; Bartnicki & Castro, 1969; van den Wijngaard et al., 1991; Poelarends et al., 1999; van Hylckama Vlieg et al., 2001). Substrates include both chlorinated and brominated C2 and C3 alcohols such as 2-chloroethanol, 1,3-dichloro-2- propanol and 2,3-dichloro-1-propanol and their brominated analogues, but also aromatic halohydrins such as 1-chloro-2-phenylethanol. The observation that the dehalogenation of chiral halohydrins can proceed with high enantioselectivity, has attracted the broad interest in these enzymes (Nakamura et al., 1992; Nakamura et al., 1994; Assis et al., 1998; Lutje Spelberg et al., 1999; Lutje Spelberg et al., 2001; van Hylckama Vlieg et al., 2001).

X OH HheC X OH O + β α R - HX R R

Figure 1. Schematic representation of the reversible reaction catalyzed by the enantioselective halohydrin dehalogenase from A. radiobacter AD1. The enzyme preferably converts one enantiomer of a racemic mixture of a vicinal halohydrin. The backward reaction, the epoxide ring opening by a nucleophile, is not only enantioselective but also highly β-regioselective.

The amino acid sequences of six halohydrin dehalogenases have been obtained so far (Yu et al., 1994; Poelarends et al., 1999; van Hylckama Vlieg et al., 2001). Based on their sequences, the six enzymes have been classified in three groups of two enzymes each, which are referred to as group A, B and C (van Hylckama Vlieg et al., 2001). Within the groups, the members share more than 80% sequence identity, whereas the pair wise sequence identity between the groups is only 25-35%. There are striking differences in enantioselectivity for the conversion of substrates by enzymes of different groups. For example, the enantioselectivity of the HheBGP1 of group B and HheC of group C is opposite to each other for aromatic halohydrins like 2-chloro-1-phenylalcohol (van Hylckama Vlieg et al., 2001). Sequence comparisons with other protein sequences showed that halohydrin dehalogenases are homologous to members of the short-chain dehydrogenase/reductase (SDR) family (Jörnvall et al., 1995; van Hylckama Vlieg et al., 2001). SDR proteins catalyze the reversible conversion of a hydroxyl group into a ketone or aldehyde group using NADP+ or NAD+ as a cofactor. Halohydrin dehalogenases lack the specific signature of a nucleotide binding sequence motif, which is in agreement with the observation that they do not need a cofactor. On the other hand, the catalytic serine and tyrosine residues of the SDR proteins are strictly conserved in halohydrin dehalogenases, although the catalytic lysine is conservatively replaced by an arginine (van Hylckama Vlieg et al., 2001). Van Hylckama Vlieg et al. have proposed a catalytic mechanism for the halohydrin dehalogenases, based on the well-studied catalytic mechanism of the SDR proteins (Jörnvall et al., 1995). The hydroxyl group of a halohydrin is thought to be hydrogen bonded to the catalytic serine and tyrosine residues. The arginine residue is proposed to increase the pKa of the tyrosine residue, thereby enabling the tyrosine to abstract a proton from the vicinal hydroxyl group of the halohydrin. The activated hydroxyl group then can perform a nucleophilic attack on the β-carbon atom, resulting in the cleavage of the carbon-halogen bond and the concurrent ring closure of the epoxide. To favor the breaking of the carbon-halogen bond, the halogen atom/halide ion is possibly stabilized by a halide binding site, as has been observed in the structures of several other types of dehalogenases (Verschueren et al., 1993; Ridder et al., 1999). The reverse reaction, the ring opening of epoxides by a nucleophile catalyzed by halohydrin dehalogenases, has also been studied. This resulted in the observation that not only chloride and bromide ions, but also azide and cyanide ions can serve as a

41 nucleophile in the reaction catalyzed by these enzymes (Nakamura et al., 1991; Lutje Spelberg et al., 2001). For some substrates, the nucleophilic ring opening of epoxides also proceeds with high enantioselectivity (Nakamura et al., 1991; Lutje Spelberg et al., 2001). Furthermore, the nucleophilic epoxide ring opening catalyzed by halohydrin dehalogenases may proceed with a different regioselectivity as compared to the chemical ring opening. It has been shown that the azidolysis of (substituted) styrene oxides by the halohydrin dehalogenase from A. radiobacter AD1 (HheC) is not only highly enantioselective, but also highly β-regioselective (Lutje Spelberg et al., 2001). These properties give this enzyme a high potential to be used as a biocatalyst in the regio- and enantioselective synthesis of chiral epoxides, haloalcohols, azidoalcohols and cyanoalcohols. We have started a crystallographic investigation of halohydrin dehalogenases to gain insight in the structural details of the catalytic machinery of these enzymes. Such knowledge may assist in the rational improvement of the catalytic properties and substrate/product preferences of these enzymes.

Crystallization, data collection and processing

A. radiobacter halohydrin dehalogenase C (HheC) was prepared as described before (van Hylckama Vlieg et al., 2001). Crystals were obtained from hanging drop experiments at room temperature using drops of 2 µl of protein and 2 µl of reservoir solution suspended over 1 ml of reservoir solution. A reservoir solution containing 50 - 70% saturated ammoniumsulfate solution (saturated at room temperature) in 100 mM bis- Tris (bis[2-hydroxyethyl]imino-tris-hydroxymethyl]methane) buffer of pH 6.5 to 7.1, resulted in the formation of multiple bipyramidal crystals of dimensions up to 0.4 x 0.3 x 0.3 mm3, grown within two weeks (Figure 2). The hanging drops initially show some protein precipitation, but this does not prevent crystal formation. Addition of 5 mM DTT to the reservoir solution prevents this precipitation, which suggests that free cysteine residues in the protein are a likely cause for this. A native dataset to 3.0 Å was collected in house at room temperature on a MacScience image plate system using Cu-Kα radiation from a rotating anode generator. The data were processed using DENZO and SCALEPACK (Otwinowski and Minor, 1997). The dataset contained 148242 measurements, which were reduced to 13874 unique reflections with an average Rmerge of 13.0% on intensities. The completeness of the data is

99.5%. The crystals belong to space group P41212 or P43212 with cell axes a = b = 104.5 Å and c = 121.4 Å. This results in a unit cell volume of 1.33 * 106 Å3. With two monomers 3 of approximately 27.5 kDa in the asymmetric unit, the volume per unit mass, Vm, is 3.01 Å Da-1 (Matthews, 1968). These values are within the range of 1.7 - 3.5 Å3 Da-1 usually found for proteins. The calculated solvent content is 59%, assuming a specific volume of 0.74 cm3/g for the protein molecule.

Figure 2. A crystal of halohydrin dehalogenase HheC from A. radiobacter AD1.

Preliminary phases for HheC crystals have been obtained by a multiple anomalous dispersion (MAD) experiment around the Br edge on HheC crystals obtained in the presence of NaBr. A full description of the procedure to obtain useful phase information is described in Chapter 4.

References

Assis HMS, Sallis PJ, Bull AT, Hardman DJ: Biochemical characterization of a haloalcohol dehalogenase from Arthrobacter erithii H10a. Enz Microb Tech 1998, 22:568-574. Bartnicki EW, Castro CE: Biodehalogenation. The pathway for transhalogenation and the stereochemistry of epoxide formation from halohydrins. Biochemistry 1969, 8, 4677-4680. Castro CE, Bartnicki EW: Biodehalogenation. Epoxiation of halohydrins, epoxide opening, and transhalogenation by a Flavobacterium sp. Biochemistry 1968, 7, 3213-3218. Fetzner S: Bacterial dehalogenation. Appl Microbiol Biotechnol 1998, 50:633-657. Fetzner S, Lingens F: Bacterial dehalogenases: biochemistry, genetics, and biotechnological applications. Microbiol Rev 1994, 58:641-685. Jörnvall H, Persson B, Krook M, Atrian S, Gonzalez Duarte R, Jeffery J, Ghosh D: Short-chain dehydrogenases/reductases (SDR). Biochemistry 1995, 34:6003-6013. Lutje Spelberg JH, van Hylckama Vlieg JET, Bosma T, Kellogg RM, Janssen DB: A tandem enzyme reaction to produce optically active halohydrins, epoxides and diols. Tetrahedron: Asymmetry 1999, 10:2863-2870. Lutje Spelberg JH, van Hylckama Vlieg JET, Tang L, Janssen DB, Kellogg RM: Highly enantioselective and regioselective biocatalytic azidolysis of aromatic epoxides. Org Lett 2001, 3:41-43. Matthews BW: Solvent content of protein crystals. J Mol Biol 1968, 33:491-497. Nakamura T, Nagasawa T, Yu F, Watanabe I, Yamada H: A new enzymatic synthesis of (R)-γ- chloro-β-hydroxynitrile. Appl Environ Microbiol 1994, 60:1297-1301. Nakamura T, Nagasawa T, Yu F, Watanabe I, Yamada H: Resolution and some properties of enzymes involved in enantioselective transformation of 1,3-dichloro-2-propanol to (R)-3- chloro-1,2-propanediol by Corynebacterium sp. strain N-1074. J Bacteriol 1992, 174:7613-7619. Nakamura T, Nagasawa T, Yu F, Watanabe I, Yamada H: A new catalytic function of halohydrin hydrogen-halide-lyase, synthesis of beta-hydroxynitriles from epoxides and cyanide. Biochem Biophys Res Commun 1991, 180:124-130. Otwinowski Z, Minor W: Processing of X-ray diffraction data collected in oscillation mode. Meth Enzymol 1997, 276:307-326. Poelarends GJ, van Hylckama Vlieg JET, Marchesi JR, Freitas Dos Santos LM, Janssen DB: Degradation of 1,2-dibromoethane by Mycobacterium sp. strain GP1. J Bacteriol 1999, 181:2050- 2058. Ridder IS, Rozeboom HJ, Kalk KH, Dijkstra BW: Crystal structures of intermediates in the dehalogenation of haloalkanoates by L-2-haloacid dehalogenase. J Biol Chem 1999, 274, 30672- 30678. van den Wijngaard AJ, Reuvekamp PT, Janssen DB: Purification and characterization of haloalcohol dehalogenase from Arthrobacter sp. strain AD2. J Bacteriol 1991, 173:124-129.

44 van Hylckama Vlieg JET, Tang L, Lutje Spelberg JH, Smilda T, Poelarends GJ, Bosma T, van Merode AEJ, Fraaije MW,Janssen DB: Halohydrin dehalogenases are structurally and mechanistically related to short-chain dehydrogenases/ reductases. J Bacteriol 2001, 183:5058-5066. Verschueren KH, Seljée F, Rozeboom HJ, Kalk KH, Dijkstra BW: Crystallographic analysis of the catalytic mechanism of haloalkane dehalogenase. Nature 1993, 363, 693-698. Yu F, Nakamura T, Mizunashi W, Watanabe I: Cloning of two halohydrin hydrogen-halide lyase genes from Corynebacterium sp. strain N-1074 and structural comparison of the genes and gene products. Biosci Biotechnol Biochem 1994, 58, 1451-1457.

Chapter 4: Structure and mechanism of a bacterial haloalcohol dehalogenase: a new variation of the short-chain dehydrogenase/reductase fold without an NAD(P)H binding site

René M. de Jong, Jan J. W. Tiesinga, Henriëtte J. Rozeboom, Kor H. Kalk, Lixia Tang, Dick B. Janssen & Bauke W. Dijkstra

Abstract

Haloalcohol dehalogenases are bacterial enzymes that catalyze the cofactor-independent dehalogenation of vicinal haloalcohols such as the genotoxic environmental pollutant 1,3-dichloro-2- propanol, thereby producing an epoxide, a chloride ion and a proton. Here we present X-ray structures of the haloalcohol dehalogenase HheC from Agrobacterium radiobacter AD1, and complexes of the enzyme with an epoxide product and chloride ion, and with a bound haloalcohol substrate mimic. These structures support a catalytic mechanism in which Tyr145 of a Ser-Tyr-Arg catalytic triad deprotonates the haloalcohol hydroxyl function to generate an intramolecular nucleophile that substitutes the vicinal halogen. Haloalcohol dehalogenases are related to the widespread family of NAD(P)H-dependent short-chain dehydrogenases/reductases (SDR family), which use a similar Ser-Tyr-Lys/Arg catalytic triad to catalyze reductive or oxidative conversions of various secondary alcohols and ketones. Our results reveal the first structural details of an SDR- related enzyme that catalyzes a substitutive dehalogenation reaction rather than a redox reaction, in which ahalide-binding site is found at the location of the NAD(P)H binding site. Structure-based sequence analysis reveals that the various haloalcohol dehalogenases have likely originated from at least two different NAD-binding SDR precursors.

Published in: EMBO Journal 2003, 22, 4933-4944.

Introduction

Dehalogenases are enzymes that are able to cleave carbon-halogen bonds (Janssen et al., 2001). Structural characterization of haloalkane dehalogenases and haloacid dehalogenases demonstrated that these hydrolytic enzymes are evolutionarily related to widespread esterase and phosphatase families (Ollis et al., 1992; Hisano et al., 1996; Ridder and Dijkstra, 1999). Haloalcohol dehalogenases, also known as halohydrin dehalogenases or halohydrin hydrogen-halide lyases, cannot be classified in these existing dehalogenase families. Instead, they show low sequence similarity to members of the short-chain dehydrogenase/reductase (SDR) family (Smilda et al., 2001; van Hylckama Vlieg et al., 2001). This family contains redox enzymes that depend on NAD(P)H, which is bound in a characteristic dinucleotide binding fold (Rossmann fold) (Rossmann et al., 1974). They have a conserved catalytic triad of Ser, Tyr and Lys/Arg residues (Jörnvall et al., 1995; Oppermann et al., 2003), which is also present in the haloalcohol dehalogenases (van Hylckama Vlieg et al., 2001). Many structures of short-chain dehydrogenases/reductases in complex with dinucleotides and substrates have revealed the structural details of the reactions catalyzed by them (Jörnvall et al., 1995; Filling et al., 2002; Oppermann et al., 2003). In addition, the structure of a dinucleotide-binding transcription factor that lacked the catalytic tyrosine and thus oxidoreductase activity showed that the SDR fold also functions in non-enzymatic activities (Stammers et al., 2001). Haloalcohol dehalogenases catalyze the intramolecular displacement of a halogen by the vicinal hydroxyl group in 1,3-dichloro-2-propanol, yielding its corresponding epoxide, a halide ion and a proton (van Hylckama Vlieg et al., 2001). 1,3-dichloro-2- propanol is a well-known environmental pollutant that has extensively been applied in industry. It is mutagenic and genotoxic to bacterial and mammalian cells due to the spontaneous formation of the reactive epoxide epichlorohydrin (Hahn et al., 1991). Together with a well-characterized epoxide hydrolase (Rink et al., 1999; Nardini et al., 1999), the haloalcohol dehalogenase constitutes an efficient degradation pathway of 1,3- dichloro-2-propanol and epichlorohydrin into non-toxic metabolites, which can be used by the bacterium as a source of carbon and energy (Figure 1). Haloalcohol dehalogenases have received increased attention due to their broad substrate specificity and enantioselectivity towards aliphatic as well as aromatic vicinal haloalcohols (van Hylckama Vlieg et al., 2001; Lutje Spelberg et al., 2002). They also

48 efficiently catalyze the back reaction, the enantio- and β-regioselective epoxide ring opening by nucleophiles like azide and cyanide anions (Nakamura et al., 1994; Lutje Spelberg et al., 2001). These properties make them promising candidates to become tools in the synthesis of enantiopure aliphatic and aromatic epoxides and haloalcohols, as well as azido-, cyano- and other β-substituted alcohols (Lutje Spelberg et al., 1999; Lutje Spelberg et al., 2002).

Figure 1. Putative degradation route of 1,3-dichloro-2-propanol in A. radiobacter AD1. The epoxide hydrolysis steps are catalyzed by epoxide hydrolase (EH), whereas the dehalogenation steps are catalyzed by the haloalcohol dehalogenase HheC.

X-ray crystallography and site-directed mutagenesis was used to provide insight into the catalytic machinery of the haloalcohol dehalogenase HheC from the soil bacterium Agrobacterium radiobacter AD1 (van Hylckama Vlieg et al., 2001) and its evolutionary relationships with short-chain dehydrogenases/reductases. Our results extend our insight into the diversity of the superfamily of SDR-related proteins with a catalytically active member that has lost its NAD(P)H binding capability. Instead, the dinucleotide-binding site has become a spacious halide-binding site that facilitates haloalcohol dehalogenation by an SN2-type substitution mechanism, rather than a redox reaction.

Results and discussion

Structure determination

Since previously studied hydrolytic dehalogenases contain a high affinity halide- binding site (Verschueren et al., 1993a; Ridder et al., 1997), we crystallized A. radiobacter haloalcohol dehalogenase HheC in the presence of bromide ions to use their anomalous signal for phasing (Dauter et al., 2001) (see Materials & Methods). Its structure was subsequently elucidated via a three-wavelength MAD experiment on the Br-edge. Sixteen bromide ions with unusually high occupancies ranging from 0.7 to 0.95 could easily be located in the data, corresponding to the single high affinity halide-binding site in each of the sixteen molecules in the asymmetric unit. A four-fold averaged 2.8 Å resolution electron density map obtained from the MAD data was used to build an initial model, which was subsequently refined against the high resolution data of a perfectly merohedrally twinned crystal. The resulting 1.8 Å resolution model contains one biologically functional tetramer of HheC per asymmetric unit, with each monomer consisting of the first 252 out of 254 amino acids, 1 bromide ion bound in the active site, and, in total, 291 water molecules. This model was the starting model for the structure elucidation of enzyme-substrate and enzyme-product complexes. A summary of the refinement and model statistics is shown in Table 1.

Overall structure of haloalcohol dehalogenase HheC

HheC is a tetramer with 222 point group symmetry that can be regarded as a dimer of dimers (Figure 2A). The different subunits of the tetramer superimpose with r.m.s.d.’s between 0.45 and 0.55 Å for 252 Cα atoms. Each subunit consists of a seven- stranded parallel β-sheet flanked on both sides by α-helices, which resembles the characteristic Rossmann fold (Figures 2B, 2C). Two monomers form a dimer in which the longest α-helices αE and αF and part of their connecting loops associate into an intermolecular anti-parallel four helix bundle. Two of these dimers make up the full tetramer through contacts of β-strand βG and α-helix αG and their connecting loops. The β-strands come together in an anti-parallel fashion and mainly interact through hydrophobic and electrostatic side chain interactions.

Data collection & statistics of MAD Peak Inflection Remote on Br- λ = 0.9190 Å λ = 0.9196 Å λ = 0.8551 Å

Space group P2 2 2 1 1 Unit cell (Å, º) a = 118.0 , b = 292.9, c = 146.4, α = β = γ = 90 No. molecules per ASU 16 No. Br- sites (halide-binding site/other 16 (14) sites) Resolution (outer shell) (Å) 25 – 2.7 (2.8 - 2.7) 25 – 2.7 (2.8 - 2.7) 25 – 2.8 (2.9 - 2.8) a Rsym (%) overall (outer shell) 5.5 (14.5) 5.8 (12.7) 6.8 (19.0) Completeness (%) overall (outer shell) 95.9 (84.5) 95.9 (85.1) 89.4 (62.0) I/σ overall (outer shell) 15.7 (4.9) 14.9 (4.9) 10.3 (2.5) Reflections total (unique) 1718858 (139737) 1726074 (139618) 1058025 (138855) Overall phasing power - / - 2.36 / - 2.25 / - (centrics:ISO/ANO) b Overall phasing power - / 0.76 3.46 / 1.28 3.41 / 0.98 (acentrics:ISO/ANO) b (centric/acentric) c 0.26 / 0.35

Data collection of complexes HheC·Br- HheC·Cl-·(R)-SO HheC·(R)-pNPAE (twinned, α = 0.48)

Space group P43 P43212 C2 No. molecules / ASU 4 2 4 Unit cell (Å, º) a = b = 103.2 a = b = 103.9 a = 146.7 c = 117.4 c = 119.4 b = 71.8 α = β = γ = 90 α = β = γ = 90 c = 97.3 α = 90 β = 92.8 γ = 90

Resolution (outer shell) (Å) 35 – 1.8 (1.9 - 1.8) 30 - 2.6 (2.7 - 2.6) 25 – 1.9 (2.0 - 1.9) a Rsym (%) overall (outer shell) 6.6 (17.0) 9.6 (41.0) 5.6 (18.5) Completeness (%) overall (outer shell) 95.2 (46.4) 96.3 (91.7) 98.7 (93.9) I/σ overall (outer shell) 23.4 (3.2) 10.8 (2.9) 23.5 (5.8) Reflections total (unique) 1561167 (63725) 297117 (20596) 1029109 (78669)

Refinement Total atoms/water 7889 / 291 3916 / 64 8630 / 898 d R/Rfree 23.3 / 30.9 (twinned) 24.6 / 29.6 16.9 / 19.6 R.m.s.d bonds (Å)/Angles (º) 0.008 / 1.51 0.007 / 1.42 0.005 / 1.37 R.m.s.d B (Å2) (main chain/side chain) 0.85 / 1.19 0.91 / 1.61 0.68 / 1.51 Ramachandran plot (%) 84.1 / 14.0 / 1.4 / 0.5 85.6 / 13.0 / 0.5 / 0.9 85.2 / 13.4 / 0.5 / (fav./all./gen. all./disall.) e 0.9 a Rsym = Σ|I - |/ΣI, where I is the observed intensity and the average intensity. b Phasing power = r.m.s. (|FX|/E), where |FX| can be the calculated isomorphous (ISO) or anomalous (ANO) structure factor and E is the residual lack of closure c = mean figure of merit over all resolution bins d R = R based on 95% of the data used in refinement. Rfree = R based on 5% of the data withheld for the cross- validation test. e The percentage of residues in the favored, allowed, generously allowed, and disallowed regions of he Ramachandran plot.

Table I. Data collection and refinement statistics.

The fold and dimerization interfaces of HheC are typical for members of the SDR family, which mainly occur as tetramers or dimers (Jörnvall et al., 1995). The C-terminal end of each HheC monomer binds the 2-fold symmetry related subunit that lies opposite in the other dimer, and contributes a tryptophan residue (Trp249) to its active site (Figures 2A, 3). Similar interactions of C-terminal residues with the active site of another subunit have been observed in structures of other members of the SDR family (Benach et al., 1998). The relevance of this interaction for HheC is not known. Sequence alignment of the available haloalcohol dehalogenases shows that some of them lack the last nine C- terminal residues, suggesting that the interaction of the C-terminus with the opposite active site is not essential.

The halide binding site of haloalcohol dehalogenase HheC

The HheC·Br- binary complex highlights the active site, which is located in a loop- rich cavity that contains the Ser132-Tyr145-Arg149 catalytic triad and a bound bromide ion (Figures 2B, 2C). Tyr145 is hydrogen-bonded to Arg149 in the next turn of helix αF. The third residue of the catalytic triad, Ser132, is located at the tip of strand βE with its hydroxyl group making hydrogen bonds with the side chain oxygen atom and the backbone NH group of Thr134. A loop starting from β-strand βF and a small loop between β-strand βA and helix αB form a spacious halide binding site. The interactions of the halide ion are more extensive and of a more hydrophobic nature than those previously found for haloalkane dehalogenases and haloacid dehalogenases (Verschueren et al., 1993a; Ridder et al., 1997). The bromide ion interacts with the peptide nitrogen atoms of Tyr177 and Leu178, with the side chains of Pro175 and Leu178, the Cα atom of Asn176, and with the aromatic rings of Phe12, Phe186 and Tyr187 (Figure 3A).

Figure 2 (right). Stereo view of the tetrameric structure of the HheC·Br- complex. The location of the active site in each monomer is indicated by the catalytic serine, tyrosine and arginine in ball-and-stick representation and the bound bromide ion as a yellow sphere. B) Stereo view of the Cα-trace of a monomer of the HheC·Br- complex. The catalytic residues Ser132, Tyr145, Arg149, Asp80 and two internal water molecules are depicted in ball-and-stick representation. The bromide ion is shown as a yellow sphere. The hydrogen-bonding pattern that extends from the catalytic base tyrosine to Asp80 at the surface of the molecule is shown as black dashed lines. C) Secondary structure topology diagram showing the location of the sequence motifs that are responsible for halide binding, substrate binding and proton relay. The corresponding sequence motifs of the SDR enzyme family are shown in parentheses.

The halide ion also interacts with a water molecule that is bound in the back of the halide-binding site (Figure 3A), where it bridges the backbone carbonyl group of Leu178 and the amide group of the cis-peptide bond between Tyr185 and Phe186. This cis-peptide is further stabilized by a hydrogen bond between the backbone carbonyl oxygen atom of Tyr185 and the peptide nitrogen atom of Phe12. Other contacts that stabilize the overall conformation of the halide-binding site are hydrogen bonds between the side chains of Asn176 and Tyr187, and between Ser180 and the NH group of Tyr187. The tyrosine hydroxyl group further provides the docking site for the indole NH function of Trp249 from the opposite 2-fold symmetry related subunit.

Product binding in haloalcohol dehalogenase HheC

The equilibrium of the dehalogenation reaction lies almost completely to the side of the epoxide (van Hylckama Vlieg et al., 2001). Therefore, to gain insight into epoxide product binding, HheC was crystallized in the presence of NaCl and a racemic mixture of styrene oxide. This resulted in a 2.6 Å resolution ternary complex of the enzyme with a chloride ion and (R)-styrene oxide ((R)-SO) (Figures 3B, 3D). In this complex, the chloride ion occupies the halide-binding site, which shows no large conformational changes compared to its bromide-bound state in the HheC·Br- complex. The aromatic ring of styrene oxide interacts with the side chains of several aromatic and aliphatic residues that line the active site cleft. A water molecule bound between Ser132 and Tyr145 in the HheC·Br- complex has been replaced by the epoxide ring of (R)-SO. The Ser132 and Tyr145 side chains make hydrogen bonds of 2.7 Å with the epoxide oxygen atom, which are arranged in an approximately tetrahedral configuration with the two CO bonds of the epoxide ring. They position the epoxide Cβ atom close to the chloride ion bound in the halide binding site, which could explain the high β-regioselectivity of nucleophilic epoxide ring opening reactions catalyzed by HheC (Lutje Spelberg et al., 2002).

Figure 3 (right). A) A detailed overview (in stereo) of the halide binding site in the HheC·Br- complex. Residues are shown in ball-and-stick representation and hydrogen bonds are indicated by dashed lines. Trp249* is the only residue that is contributed by a opposite monomer in the tetramer. B – C) Detailed schematic representations of the active site of respectively the HheC·Cl-:(R)-SO (B) and HheC·(R)-pNPAE complex (C). Portions of a 2Fo-Fc simulated annealing electron density omit map are shown (Brünger et al., 1997), contoured at 1σ and covering the bound molecules. D – E) Simplified schematic drawing of the bound epoxide and azidoalcohol molecule in respectively the HheC·Cl-·(R)-SO complex (D) and HheC·(R)-pNPAE complex (E).

Binding of a substrate mimic in haloalcohol dehalogenase HheC

Aromatic azidoalcohols or cyanoalcohols, the products of the epoxide ring opening reaction catalyzed by HheC in vitro, can be used to study the binding of haloalcohol substrates. They can be regarded as mimics of haloalcohols, as their azide and cyanide function have similar binding properties as halides (Norne et al., 1975; Verschueren et al., 1993b). The equilibrium of the nucleophilic ring opening of an epoxide by azide lies completely at the side of the azidoalcohol product (Lutje Spelberg et al., 2002). Therefore, cocrystallization of HheC in the presence of (R)-1-para-nitro-phenyl-2- azido-ethanol ((R)-pNPAE) resulted in a stable HheC·azidoalcohol complex at 1.9 Å resolution (Figures 3C, 3E). The azide group of the compound is bound in the halide binding site of the enzyme, making similar contacts with the backbone amide groups, aromatic amino acid side chains and the structural water molecule as observed for bound halides. The secondary hydroxyl moiety of the chiral Cα atom of the azidoalcohol is bound to Ser132 and Tyr145 by hydrogen bonds of around 2.6 Å. The epoxide and azidoalcohol complexes superimpose with a very low r.m.s.d. of only 0.20 Å for 1902 atoms of a HheC monomer. The conformations of the active site residues are virtually the same, showing that the substrate mimic and product are equally well accommodated by the active site. This is in agreement with HheC being able to catalyze both the back and forward reaction. The non-specific binding mode of the epoxide and azidoalcohol side chain in the complexes could explain the broad acceptance of the enzyme of substituted aliphatic and aromatic haloalcohol and epoxide side chains (van Hylckama Vlieg et al., 2001).

Structural relationships between haloalcohol dehalogenases and short- chain dehydrogenases/reductases

A structural similarity search with HheC against the PDB database using the SCOP server (Murzin et al., 1995) yielded 20 SDR structures. The four highest scoring structures were 7α-hydroxysteroid dehydrogenase from Escherichia coli (PDB-code 1FMC, Tanaka et al., 1996), trihydroxynaphtalene reductase from Magnaporthe grisea (PDB-code 1YBV, Andersson et al., 1996) and the plant tropinone reductases I and II from Datura stramonium (PDB-code 1AE1, Nakajima et al., 1998; PDB-code 2AE1, Yamashita et al., 1999). These four enzymes are all NAD(P)H-dependent enzymes of approximately 250 residues that catalyze the dehydrogenation of a hydroxyl group or the reduction of a carbonyl group. All four structures superimpose on the structure of the

HheC·Cl-·(R)-SO complex with r.m.s. differences between 1.15 and 1.5 Å for approximately 190 Cα atoms, whereas their amino acid sequences share only 20-25% sequence identity. High structural similarity despite low sequence identity has previously been recognized as a general property of the SDR family (Jörnvall et al., 1995). A structure-based sequence alignment shows that the Ser-Tyr-Arg/Lys catalytic triads are strictly conserved, whereas two regions show high divergence between HheC and related dehalogenases on the one hand and the four most similar SDR enzymes on the other (Figure 4). The first divergent region lies between residues 175 and 189 of HheC, which corresponds to the loop between strand βF and helix αFG2 that participates in cofactor and substrate binding in the SDR enzymes. In HheC, this loop forms most of the halide-binding site (Figure 5A). In the SDR enzymes, the amino group of the carboxylamine-pyridine function of the nicotinamide adenine dinucleotide (NADH) cofactor is stabilized by a mostly conserved threonine (Thr194 of 7α-HSD) (Filling et al., 2002), whereas the structurally equivalent Ser180 in HheC stabilizes the backbone conformation of the halide-binding loop. This residue is not conserved in the other haloalcohol dehalogenases (Figure 4), which suggests that Ser180 plays no essential role in the dehalogenases. However, the corresponding loop contains the cis-peptide between Phe185 and Tyr186 that binds the halide-bound water molecule. These residues are conserved in other haloalcohol dehalogenase sequences, which may indicate a general stabilizing role for the cis-peptide in the architecture of the halide-binding site. The second and most striking difference is found in the loop that connects βA and αB and that contains the highly conserved nucleotide-binding Gly-X-Gly-X3-Gly motif of the SDR enzymes (Jörnvall et al., 1995). In HheC this motif is replaced by a NVKHFGG pattern (residues 8-14), and its larger side chains occupy the space where in the SDR enzymes part of the NADH cofactor is bound (Figures 4, 5B). The glycine-replacing Phe12 makes important backbone-backbone interactions with the cis-peptide between Phe185 and Tyr186. Phe12 has also stacking interactions with the aromatic side chain of Phe185. These interactions cause the two described loops that each interact with the cofactor in SDR enzymes, to pack closely together and form the spacious halide-binding site in HheC.

Figure 5. A) Stereo view of a superposition of a Cα trace of the two divergent loops of the HheC·Cl- ·(R)-SO complex and the ternary complex of 7α-hydroxysteroid dehydrogenase with NADH and 7- oxoglycochenodeoxycholic acid (7-GCDCA). The chloride ion and a water molecule are shown as spheres. B) Stereo view of a superposition of the active site regions of the HheC·Cl-·(R)-SO complex and the ternary complex of 7α-hydroxysteroid dehydrogenase with NADH and 7-GCDCA. Residues, substrates and cofactor are shown in ball-and-stick representation and hydrogen bonds are indicated by dashed lines. C) Representation of the proposed proton relay pathway in the HheC·Cl-·(R)-SO complex from the catalytic base Tyr145 to a water molecule (W3) in the first solvation shell of the HheC tetramer. Amino acids and epoxide substrate are shown in ball-and-stick representation. A chloride ion and water molecules are shown as spheres. D) A detailed schematic representation of the proton relay pathway in the ternary complex of 7α-hydroxysteroid dehydrogenase with NADH and 7-GCDCA.

Figure 4 (left). Sequence alignment of the haloalcohol dehalogenases HalB (Agrobacterium tumefaciens, unpublished, acc. no. AAD34609), HheAAD2 (Arthrobacter sp. AD2, van Hylckama-Vlieg et al., 2001), HheA (Corynebacterium sp. N-1074, Yu et al., 1994), HheBGP1 (Mycobacterium sp. GP1, Poelarends et al., 1999) and HheB (Corynebacterium sp. strain N-1074, Yu et al., 1994), combined with a 3D-structure-based alignment of HheC (A. radiobacter AD1, van Hylckama-Vlieg et al., 2001), and the four homologous SDR structures with the highest structural similarity to HheC (7α- hydroxysteroid dehydrogenase from E. coli, PDB-code 1FMC, Tanaka et al., 1996; trihydroxynaphtalene reductase from M. grisea, 1YBV, Andersson et al., 1996; tropinone reductases I and II from D. stramonium, 1AE1, Nakajima et al., 1998 & 2AE1, Yamashita et al., 1999). The colors correspond to those in Figure 2C. The underlined amino acids correspond to the conserved asparagine in SDR enzymes (Asn118 in 7α-HSD) (Filling et al., 2002). The boxes indicate the possible positions of acidic and basic residues that respectively define the NADH or NADPH cofactor specificity in the SDR family (Persson et al., 2003).

Catalytic mechanism of haloalcohol dehalogenases

The structural correspondence of HheC to the SDR suggests a key role for the conserved catalytic residues in the dehalogenase mechanism. The epoxide and azidoalcohol complexes of HheC show that the Ser132 and Tyr145 side chains make hydrogen bonds with the oxygen atom of the epoxide product and with the hydroxyl group of a haloalcohol substrate (Figure 6A). The direct interaction of Arg149 with Tyr145 in the otherwise hydrophobic active site cavity likely polarizes the tyrosine hydroxyl group and lowers its pKa. Deprotonation of the tyrosine would allow this residue to act as the catalytic base that accepts a proton from the hydroxyl group of a vicinal haloalcohol substrate to generate an intramolecular nucleophile that concertedly substitutes the halogen. Ser132, Tyr145 and Arg149 were previously shown to be important for the catalytic mechanism, as a Ser132Ala, Tyr145Phe and a Arg149Asn mutant are more than 10,000-fold less active than the wild type enzyme (Table 2) (van Hylckama Vlieg et al., 2001). In short-chain dehydrogenases, the catalytic tyrosine deprotonates the hydroxyl group of a substrate, and transfers the proton to the catalytic lysine via the oxygen atom of a ribose hydroxyl group of the cofactor (Figures 5D, 6B) (Liao et al., 2001, Filling et al., 2002). In HheC, the backbone carbonyl group of Asp80 is at the position of the ribose hydroxyl group, which precludes proton transfer from the substrate to the equivalent Arg149 via this route. Instead, Arg149 directly contacts the tyrosine hydroxyl group and thus can directly take up a proton from the tyrosine. Mutation of this arginine into a lysine yields an enzyme that is 400-fold less active than the wild-type enzyme (Table 2) (van Hylckama Vlieg et al., 2001). Impaired proton transfer between the tyrosine and the lysine could provide a plausible explanation for this, although the possibility of an effect on the pKa of Tyr145 or on the integrity of the active site can not be excluded.

-1 -1 -1 Enzyme Km (mM) kcat (s ) kcat/Km (s M ) wild type1 0.009 ± 0.003 22 ± 1 2.4 x 106 Ser132Ala2 -4 -4 -4 Tyr145Phe2 0.13 0.072 5.5 x 102 Arg149Lys2 0.056 0.33 5.9 x 103 Arg149Asn2 >0.40 >0.15 3.8 x 102 Asp80Asn3 0.02 ± 0.002 0.1 ± 0.002 5.0 x 103 Asp80Ala3 -4 -4 -4

1 Data taken from Tang et al., 2003. 2 Data taken from van Hylckama Vlieg et al., 2001. 3 This study. 4 No activity detectable.

Table 2. Steady-state kinetic parameters of wild-type and mutant HheC with the substrate para-nitro- 2-bromo-1-phenylethanol.

Figure 6. A) A schematic representation of the proposed catalytic mechanism of dehalogenation in HheC. The catalytic base Tyr145 activates the hydroxyl group adjacent to the halogen atom for nucleophilic attack on the halogen-bearing carbon atom. The proton is released to the solvent via Arg149, the water molecules, and Asp80. B) A schematic representation of the proposed catalytic mechanism of SDR enzymes (Filling et al., 2002, Oppermann et al., 2003). The secondary hydroxyl group is deprotonated by the catalytic tyrosine residue, with concurrent transfer of the hydride ion to NAD+. Proton release from the catalytic tyrosine is proposed to take place via the hydroxyl group of the cofactor ribose moiety, the conserved lysine amine function, and water molecules bound in the core of the enzyme.

In the short-chain dehydrogenases the catalytic lysine further interacts with one of two buried water molecules that is bound to the backbone carbonyl oxygen atom of a conserved asparagine, located at a kink in helix αF (Asn118 in 7α-HSD). The side chain of the asparagine is, in turn, stabilized by an interaction with the conserved NNAG motif (residues 94 – 97 in 7α-HSD). These water molecules are present in the majority of structurally characterized SDR enzymes and have been proposed to mediate proton relay to the solvent (Figures 5D, 6B) (Benach et al., 1998; Filling et al., 2002). In HheC, a similar water-filled enclosure is present near a kink in helix αF at Leu104, which aligns with Asn118 of 7α-HSD (Figure 5C). The bound water molecules form a continuous pattern of hydrogen bonds from Arg149 to the partly solvent exposed side chain of Asp80. This aspartate is part of a SNDX pattern (residues 78 - 81) that superimposes on the NNAG motif of the SDR family. This suggests that the side chain of Asp80 could play a critical role in proton relay from Arg149 to the solvent. Strikingly, the pH optima of the dehalogenation and the epoxide ring opening activities are significantly different, pH 8 - 9 and 4 – 5, respectively. During a dehalogenation reaction a proton needs to be removed, which would be easiest at physiological pH or higher, where the aspartate is charged. In contrast, for epoxide ring opening reactions protons are needed, and their provision would benefit from an acidic pH, where Asp80 is mostly protonated. Therefore, the observed proton relay system offers a simple explanation for the different pH optima of the dehalogenation and epoxide ring opening reactions catalyzed by HheC. Strikingly, when Asp80 is mutated into an alanine no activity can be detected (Table 2). Moreover, an

Asp80Asn mutant of HheC has a 220-fold reduced kcat for the dehalogenation reaction with respect to the wild-type enzyme, whereas the Km is approximately the same. The similar reduction in kcat of the Asp80Asn and Arg149Lys mutant further supports the importance of the arginine and aspartate for catalysis.

On the origin of haloalcohol dehalogenase activity

The structure of HheC reveals that its fold resembles that of members of the NAD(P)H-dependent SDR family. Together with the clear homology of HheC to the SDR enzymes (20 - 25 % identity; van Hylckama Vlieg et al., 2001) this evidences a divergent evolutionary relationship. The SDR family is characterized by conserved sequence motifs for the catalytic residues, the NAD(P)H-binding site, and the proton relay system (Filling et al., 2002, Oppermann et al., 2003). HheC has only retained the catalytic residues,

62 whereas the sequence segments that correspond to the SDR cofactor-binding site and NNAG motifs define the halide-binding site and proton relay system in HheC. However, these sequences are not strictly conserved in other haloalcohol dehalogenases, which suggests diversity in halide binding and proton relay. The SDR family is of old origin and widespread in all kingdoms of life, with e.g. more than 60 different enzymes in humans and up to 46 copies in the sequenced genomes of various microbial species (Jörnvall et al., 1999, Kallberg et al., 2001). In contrast, only six haloalcohol dehalogenase genes have been isolated from selected strains of soil bacteria that degrade 1,3-dichloro-2-propanol and 1,2-dibromoethane (van Hylckama Vlieg et al., 2001). Whereas haloalkane dehalogenases are found in the genomes of several microbial species (Jesenská et al., 2000), a BLAST search (Altschul et al., 1997) with the haloalcohol dehalogenase sequences against microbial genomes did not find any haloalcohol dehalogenase sequence, but only retrieved sequences of 15-35% identity that contain the characteristic SDR NAD(P)H-binding Gly-rich motif. The rare and exclusive occurrence of haloalcohol dehalogenases in pollutant-degrading soil bacteria raises questions on their evolutionary origin. SDR enzymes have been grouped into seven different coenzyme-based subfamilies (Kallberg et al., 2002, Persson et al., 2003) on the basis of the nature of the residues at positions 32, 33, 34, and 15 (HheC numbering). The C-type (HheC and HalB) and A-type (HheA and HheAAD2) haloalcohol dehalogenases contain an Asp at position 32, suggesting that they have evolved from an NAD-binding precursor, rather than an

NADP-dependent enzyme (Figure 4). Intriguingly, HheB and HheBGP1 have the characteristics of a different NAD-binding subfamily (with an Asp at position 34), suggesting that these latter enzymes have originated from a different precursor than the A- and C-type dehalogenases. Furthermore, the B-type dehalogenases (HheB and

HheBGP1) share only 24% sequence identity with the A and C-type dehalogenases, which themselves are 33% identical, and, as a major difference, they lack the sequence segment that corresponds to β-strand βC and α-helix αD in HheC (Figure 4). This deletion in the Rossmann fold (Rossmann et al., 1974) has previously been overlooked as a result of misalignment of the sequences (Yu et al., 1994; Smilda et al., 2001; van Hylckama Vlieg et al., 2001). Unfortunately, it is impossible to reconstruct the evolutionary history of the isolated haloalcohol dehalogenases without knowledge of their specific precursor enzymes. It is conceivable, however, that an NAD-dependent SDR precursor could have

63 accommodated the halogen of a haloalcohol substrate in the apolar nicotinamide binding site of the cofactor and used the conserved Ser-Tyr-Lys/Arg catalytic triad to promote the substitution of the halogen. Some SDR enzymes could thus fortuitously have favoured a rudimentary haloalcohol dehalogenating activity next to an NAD-dependent redox activity. Evidence is accumulating that alternative enzymatic reactions, distinct from the normally catalyzed ones, may play an important role in the diversification of enzymes (see e.g. O'Brien and Herschlag, 1999). The recruitment of NAD-binding SDR enzymes for such a haloalcohol dehalogenase activity may have occurred only very recently in response to elevated concentrations of halogenated pollutants in the environment. Although it can be argued that the low sequence identities of haloalcohol dehalogenases to the currently known bacterial SDR sequences (15-35%) may suggest that they diverged a long time ago, the enormous diversity of soil bacteria in the environment (Torsvik and Øvreås, 2002) leaves the possibility open that more closely related SDR enzymes exist, which have not been isolated so far. It might be possible to test whether a limited number of mutations can convert an SDR enzyme into a dehalogenase in the laboratory by directed evolution experiments that select for haloalcohol dehalogenating activity.

Material and methods

Crystallization of HheC

Haloalcohol dehalogenase HheC was purified as published elsewhere (van Hylckama Vlieg et al., 2001). Crystals of HheC were prepared as described before (de Jong et al., 2002). Bromide-derivatized crystals were prepared by cocrystallization. NaBr was added to a final concentration of 50 mM to the standard reservoir solution. Hanging drops were prepared of 2 µL of a protein solution of 7.5 mg/mL and 2 µL of the reservoir solution. Crystals grew in a few days at room temperature. Crystals of a HheC·Cl-·(R)-SO complex were also obtained by cocrystallization. Sulphuric acid was used to prepare dialysis and crystallization buffers to ensure that no halides were present in the solutions. A racemic mixture of (R)- and (S)-styrene oxide solubilized in a small amount of dimethylsulfoxide (DMSO) (< 1 µL) was added to 50 µL of the purified and dialyzed protein solution (7.5 mg/mL) to a final concentration of 20 mM. A small volume (1 µL) of a concentrated NaCl solution was added to a final concentration of 10 mM. After equilibrating the protein solution for 30 minutes, non-solubilized SO was

64 spun down at 3000 g to prevent solid material to be transferred to the crystallization medium. Hanging drops of a total volume of 3 µL were prepared by mixing 1.5 µL of the saturated protein-substrate mixture with 1.5 µL of the reservoir solution described above.

Bipyramidal crystals in the tetragonal space group P43212 appeared in a few days. A HheC·azidoalcohol complex was obtained by following the same solubilization and crystallization procedure as described above for the HheC·Cl-·(R)-SO complex, using a final concentration of 25 mM of enantiopure (R)-1-para-nitro-phenyl-2-azido-ethanol ((R)- pNPAE) (97% enantiomeric excess). Plate-like crystals in space group C2 grew in 3 - 5 days. In all cases a cryo solution was made by replacing the water in the well solution by a 20% glycerol solution.

Structure determination and refinement

A crystal of space group P21212 (cell dimensions a = 117.5 Å, b = 293.1 Å, and c = 146.5 Å) containing four HheC tetramers per asymmetric unit, was used in a three- wavelength anomalous dispersion experiment collected on a MAR CCD detector at the ESRF synchrotron beam line BM14. The wavelength of the X-rays was tuned around the absorption K-edge of bromide deduced from an edge-scan of the crystal. Wavelengths of the peak and inflection point dataset were set to 0.9190 Å and 0.9196 Å respectively. Data was collected to 2.8 Å resolution over a 90 degree rotation of the crystal with an interval of 0.2 degrees. 30 Frames of the peak and inflection point datasets were recollected using an interval of 0.1 degrees due to severe overlaps in this region of the data. A remote wavelength dataset (λ = 0.8551 Å) was collected over 90 degrees with a 0.2 degree interval. All data were processed using DENZO and SCALEPACK (Table 1) (Otwinowski and Minor, 1997). The resulting scaled anomalous intensities collected at the peak wavelength were used to search for bromide positions with the program Shake-and- Bake (Weeks and Miller, 1999). Coordinates and occupancies of the bromide sites were subsequently refined against the data of all three wavelengths with the program SHARP (Table 1) (De la Fortelle and Bricogne, 1997). The positions of the 16 highest occupied sites per asymmetric unit formed approximately tetrahedral clusters of 4 atoms when expanded over the whole unit cell, suggesting that they corresponded to 16 bromide ions bound to the halide-binding sites of 16 HheC monomers. These 16 sites were subsequently used to find the NCS operators between the corresponding tetrameric HheC molecules using FINDNCS (CCP4, 1994). Three of the four NCS operators were refined in an averaging procedure using DM (Cowtan, 1994), using a combination of averaging,

65 solvent flattening and histogram matching. A spherical mask with a radius of 35 Å around the center of gravity of the 4-atom clusters made with the program NCSMASK (CCP4, 1994) was used in these calculations. The resulting three-fold averaged electron density maps were of high quality and allowed the manual building of 90% of the residues of a HheC monomer. This initial model was subsequently used as a search model for a molecular replacement experiment using data of a near-perfectly merohedrally twinned crystal of

HheC in space group P43 (as indicated by TRUNCATE (CCP4, 1994) and Todd Yeates’ Twinning Server (Yeates, 1997)). The data was collected at the Elettra synchrotron in Triëste (Italy) to 1.8 Å resolution at 100 K using an X-ray wavelength of 0.93 Å over a 180 degree rotation of the crystal with an interval of 0.5 degrees. The twin-related reflections (twinning fraction α = 0.48) were averaged with the program SFTOOLS (CCP4, 1994). Four HheC monomers in each twin domain were found by molecular replacement in AMORE (Navaza et al., 1997). One twin domain was subsequently refined using twinning scripts available in the program suite CNX (Accelrys Inc., San Diego). Diffraction data of crystals of the HheC·Cl-·(R)-SO complex (2.6 Å resolution) and the HheC·(R)-pNPAE complex (1.9 Å resolution) were collected at the ID14-2 beam line at the ESRF in Grenoble (Table 1). All data from crystals of the complexes were collected at 100 K using an X-ray wavelength of 0.93 Å. Initial phases for these data sets were obtained either from rigid body fitting in CNX or, in the case of the new crystal form, by molecular replacement calculations in AMORE (Navaza et al., 1997). Model coordinates for the substrate molecules were obtained by building and minimizing them in the program QUANTA (Accelrys Inc., San Diego). Parameter and topology files for these compounds were obtained from the Hic-Up Server (Kleywegt and Jones, 1998). All data processing was carried out using DENZO and SCALEPACK (Otwinowski and Minor, 1997). All refinements were carried out using the program suite CNX (Accelrys Inc., San Diego). All manual building of protein structures was carried out in QUANTA (Accelrys Inc., San Diego) and Xtalview (McRee, 1999). Figures were prepared using MOLSCRIPT (Kraulis, 1991) and RASTER3D (Merrit and Bacon, 1997).

Site-directed mutagenesis and kinetic characterization

Site-directed mutagenesis, purification of mutant enzymes and the measurement of the activity of the Asp80Asn mutant were routinely performed as described before (van Hylckama Vlieg et al., 2001).

Structural alignment and sequence analysis

Iterative structural alignment of HheC and the four most similar crystal structures of SDR enzymes and the subsequent sequence alignment of other available haloalcohol dehalogenase sequences on the structural alignment were performed with the program SEQUOIA (Bruns et al., 1999).

References

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Materials and methods

Crystallization

Haloalcohol dehalogenase HheC was purified and crystallized as described before (van Hylckama Vlieg et al., 2001; de Jong et al., 2002, 2003). Sulfuric acid was used to prepare crystallization buffers to ensure that no halide ions were present in the crystallization solutions. In all crystallization experiments, the protein concentration of the stock solution was 7.5 mg/mL.

Crystals of a HheC:H2O:(R)-pNSO complex were obtained by cocrystallization. A solution of (R)-pNSO (e.e. 97.5%) solubilized in a small amount of dimethylsulfoxide (< 1 µL) was added to 50 µL of the purified and dialyzed protein solution to a final concentration of 10 mM. A small volume (1 µL) of a concentrated NaOCN solution was added to a final concentration of 5 mM. After equilibrating the protein solution for 30 minutes, non-solubilized (R)-pNSO was spun down at 3000 g to prevent solid material to be transferred to the crystallization solution. Hanging drops of a total volume of 3 µL were prepared by mixing 1.5 µL of the protein-substrate mixture with 1.5 µL of the standard reservoir solution. Plate-shaped crystals with space group C2 appeared in a few days, and diffracted to 1.7 Å. Each asymmetric unit contains four HheC monomers.

A HheC:H2O:(S)-styrene oxide (SO) complex was obtained by following the same solubilization and crystallization procedure as described above for the

HheC:H2O:(R)-pNSO complex, using a final concentration of 10 mM of enantiopure (S)- SO (e.e. = 97.5%). Additionally, a catalytic amount of nitrite (0.01 mM) was added to the concentrated enzyme solution to enzymatically convert any remaining (R)-SO impurities from the (S)-SO solution. After 30 minutes incubation, 5 mM NaCl was added to the solution and crystallization drops were set up. Bipyramidal crystals with space group

P43212 grew in a few days. The crystals contained two HheC monomers in the asymmetric unit, and diffracted to 2.7 Å.

A HheC:H2O:(S)-pNSO complex was obtained by the same protocol as described for the HheC:H2O:(R)-pNSO complex, using a final concentration of 10 mM of enantiopure (S)-pNSO (e.e. = 97.5%). Bipyramidal crystals appeared in a few days, and diffracted to 1.9 Å. They had space group P21212, and contained sixteen HheC monomers in the asymmetric unit.

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Chapter 5: Structural basis for the enantioselectivity of epoxide ring opening reactions catalyzed by haloalcohol dehalogenase HheC

René M. de Jong, Jan J. W. Tiesinga, Allesandra Villa, Lixia Tang, Dick B. Janssen & Bauke W. Dijkstra

Abstract Haloalcohol dehalogenase HheC catalyzes the enantioselective dehalogenation of vicinal haloalcohols to epoxides, as well as the enantioselective and β-regioselective nucleophilic ring opening of epoxides by azide. X-ray structures of HheC in complex with bound dehalogenation products and a haloalcohol mimic previously showed that catalysis depends on a halide-binding site and a Ser-Tyr-Arg catalytic triad (chapter 4). Here we present X-ray structures of complexes of HheC with the favored and unfavored enantiomers of para-nitro-styrene oxide (R)-pNSO and (S)-pNSO with a water molecule, rather than a halide ion, occupying the halide-binding site. The oxygen atom of the favored (R)-enantiomer of pNSO is hydrogen bonded to the oxygen atoms of catalytic residues Ser132 and Tyr145. The unfavored (S)- enantiomer instead binds in a non-productive ‘upside-down’ manner, directing its Cβ atom of the epoxide ring towards the catalytic residues, whereas the oxygen atom makes a hydrogen bond with the water in the halide-binding site. Comparison of the two complexes and calculation of the relative Gibbs binding energy suggest that the non-productive binding of the unfavored (S)-enantiomer forms the basis of the enantioselectivity of the epoxide ring opening of aromatic epoxides by azide catalyzed by HheC. The results indicate that the enantioselectivity of the nucleophilic ring opening reaction of aromatic epoxides by azide not only depends on the the lower affinity of HheC for (S)- enantiomers, but also for a great extend on the low kcat due to their inability to obtain a reactive orientation in the active site.

To be published.

Introduction

Enantiopure molecular building blocks are important for the synthesis of various biologically active compounds, creating a demand for enantioselective catalysts to produce them. Enzymes are perfectly suited to catalyse enantioselective conversions, because of the chirality of their amino acid building blocks (Feringa and van Delden, 1999). Moreover, they offer green synthesis routes that proceed under mild conditions. Several classes of enzymes, like lipases and esterases, have been optimised for specific enantioselective conversions (Jaeger & Reetz, 1998; Bornscheuer, 2002). Furthermore, efforts are made to discover new enzymes that could give access to currently unavailable enantiopure compounds (Marrs et al., 1999). Haloalcohol dehalogenase HheC from Agrobacterium radiobacter AD1 is such an potentially useful enzyme (van Hylckama Vlieg et al., 2001, Janssen and de Vries, 2003). It enantioselectively dehalogenates various aliphatic and aromatic vicinal halohaloalcohols, like for example 1,3-dichloro-2-propanol and 2-phenyl-1-chloro-2- ethanol, producing HCl and the corresponding epoxides epichlorohydrin and styrene oxide. X-ray structures of HheC in complex with a haloalcohol mimic and with dehalogenation products have provided structural insights into its catalytic mechanism (Figure 1A) (de Jong et al., 2003). The hydroxyl group of the vicinal haloalcohol substrate interacts with the serine and tyrosine of a Ser132-Tyr145-Arg149 catalytic triad (Figure 1C), whereas the halogen is bound in a halide-binding site

Figure 1 (right). A) The tetrameric structure of haloalcohol dehalogenase (de Jong et al., 2003). The Ser-Tyr-Arg catalytic triad of each monomeric subunit is shown in ball-and-stick representation. A bromide ion that is bound in the halide-binding site is shown as a sphere. B) A detailed overview of the hydrophobic halide-binding site with a bound bromide ion (de Jong et al., 2003). Trp249* is the only residue that is contributed by an opposite monomer in the tetramer. C) An overview of the active site of the HheC:H2O:(R)-pNSO complex. The oxygen atom of the epoxide ring of (R)-pNSO is hydrogen bonded to Ser132 and Tyr145 of the catalytic triad. Arg149 is hydrogen bonded to Tyr145, and forms a proton relay system with two buried water molecules and the side chain of the partly solvent exposed Asp80 (de Jong et al., 2003). The halide-binding site is occupied by water molecule, instead of a halide ion. Portions of a 2Fo-Fc simulated annealing electron density omit map are shown (Brünger et al., 1997), contoured at 1σ and covering the bound (R)-pNSO and water molecule. Residues are shown in ball-and-stick representation and hydrogen bonds are indicated by black dashed lines. D) A schematic representation of the catalytic mechanism of the dehalogenation reaction catalyzed by HheC. The catalytic base tyrosine takes up a proton from the hydroxyl group of the aromatic haloalcohol, which concurrently substitutes the halogen, thereby producing the corresponding epoxide (R)-SO and a chloride ion. E) A schematic representation of the catalytic mechanism of the epoxide ring opening reaction catalyzed by HheC. At low pH, the tyrosine donates a proton to the oxygen atom of the epoxide ring of (R)-SO, with the concurrent attack of a bound nucleophile on its Cβ carbon atom, thereby producing the corresponding (R)-azidoalcohol.

(Figure 1B). The hydroxyl oxygen, activated by the catalytic base Tyr145, substitutes the halogen in an SN2-type intramolecular substitution reaction (Figure 1D). The pKa of the tyrosine base is decreased by Arg149, which also relays the abstracted proton via two buried water molecules to the partly solvent-exposed Asp80. Interestingly, the halide-binding site of HheC can also accommodate other small negatively charged ions like azide, cyanide and nitrite, which can replace the halide ion in the reverse reaction, the enantioselective and β-regioselective nucleophilic ring opening of epoxides (Figure 1E) (Lutje Spelberg et al., 2001, 2002). The ring opening reaction requires acidic conditions (pH 4-5), which allow the tyrosine to protonate a bound epoxide substrate, while an anionic nucleophile attacks the Cβ atom of the epoxide ring, producing the corresponding β-substituted alcohol. These properties make HheC a promising biocatalyst to produce various enantiopure epoxides and β-substituted alcohols (Janssen & de Vries, 2003). Intriguingly, the enantioselectivity of the dehalogenation reaction of aromatic haloalcohols is much lower than the corresponding reverse ring opening reaction of aromatic epoxides. The dehalogenation of racemic para-nitro-2-bromo-1-phenylethanol (pNSHH) results in a rapid conversion of the (R)-enantiomer, but a much slower conversion of the (S)-enantiomer (E = 150) (Lutje Spelberg et al., 2002; Tang et al., 2003). Kinetic analysis of this reaction showed that the enantioselectivity depends on the weaker binding and the slower reactivity of (S)-pNSHH (Tang et al., 2003). In contrast, the nucleophilic ring opening reaction of racemic para-nitro-styreneoxide (pNSO) by azide results in a very rapid conversion of (R)-pNSO to (R)-para-nitro-2-azido-1-phenylethanol ((R)-pNSAA), whereas conversion of the (S)-enantiomer by the enzyme cannot be detected at all (E > 200). Unfortunately, a detailed kinetic analysis of this latter reaction is not possible due to the low solubility of the epoxide in water (<2 mM). Here we present evidence from high resolution X-ray data that both (R)- and (S)- pNSO bind to HheC in a nearly identical way, but with the crucial difference that (S)-pNSO adopts a non-productive binding mode. Thermodynamic integration calculations further indicate that the Gibbs energy difference between the observed (R) and (S) bound states are very low, suggesting comparable binding affinities of HheC for the different enantiomers. These results indicate that the outstanding enantioselectivity of HheC in nucleophilic ring opening reactions of aromatic epoxides by azide is caused by the non- productive binding of the unfavored (S)-enantiomer.

Data statistics HheC:H2O:(R)-pNSO HheC:H2O:(S)-pNSO

Space group C2 P212121

No. tetramers / ASU (chains / ASU) 1 (4) 4 (16)

Unit cell (Å, º) a = 146.4 a = 118.8 b = 72.0 b = 293.6 c = 97.4 c = 146.7 α = 90 α = β = γ = 90 β = 92.9 γ = 90

Resolution (Å) 35 – 1.7 35 - 1.9 a Rsym (%) overall (outer shell) 9.6 (47.7) 8.0 (25.8) Completeness (%) overall (outer shell) 99.1 (95.6) 90.7 (92.3) I/σ overall (outer shell) 5.9 (2.2) 14.5 (3.5) Reflections total (unique) 890863 (110688) 5753736 (313025)

Refinement

Total atoms / water 8630 / 895 28976 / 3298 b R/Rfree 20.2 / 23.0 21.0 / 24.2 R.m.s.d. bond (Å) / Angles (º) 0.004 / 1.32 0.006 / 1.36 R.m.s.d. B (Å2) (main chain/side chain) 0.89 / 1.34 0.95 / 1.59

Ramachandran plot (%)

Favored 85.9 85.9 Allowed 12.8 12.1 Generously allowed 1.3 2.0 Disallowed 0.0 0.0 a Rsym = Σ|I - |/ΣI, where I is the observed intensity and the average intensity. b R = R based on 95% of the data used in refinement. Rfree = R based on 5% of the data withheld for the cross-validation test.

Table I. Data collection and refinement statistics.

Data collection and refinement

Diffraction data of crystals of the complexes were collected at 100 K using an X- ray wavelength of 0.93 Å at the ID14-1 and ID14-2 beam lines of the ESRF in Grenoble. All data processing was carried out using DENZO and SCALEPACK (Otwinowski & Minor, 1997), while refinements were done using the program suite CNX (Accelrys Inc., San Diego). Initial phases for these data sets were obtained from rigid body fitting in CNX (Accelrys Inc., San Diego). Model coordinates for the substrate molecules were obtained by building and energy-minimizing them in the program QUANTA (Accelrys Inc., San Diego). Parameter and topology files for these compounds were obtained from the Hic-Up Server (Kleywegt & Jones, 1998). When needed, the protein models were manually rebuilt in QUANTA (Accelrys Inc., San Diego) and Xtalview (McRee, 1999). Data collection and refinement statistics are summarized in table I. Figures of molecular models were prepared using MOLSCRIPT (Kraulis, 1991) and RASTER3D (Merritt & Bacon, 1997), unless otherwise stated.

Computational techniques

The obtained structure of the HheC:H2O:(R)-pNSO complex was used as the starting structure for the molecular dynamics calculations. The bound epoxide in one of the four active sites of the tetrameric enzyme was retained, whereas the other three epoxides were replaced by water molecules. The tetrameric enzyme and the single bound epoxide were hydrated in a box containing ~27,000 simple point charge (SPC; Berendsen et al., 1981) water molecules. Sixteen Na+ ions were added to balance the total charge of the system. The enzyme and epoxide were described using the GROMOS96 (43a2) force field (van Gunsteren et al., 1996; Schuler & van Gunsteren, 2000), in which aliphatic hydrogen atoms are treated as united atoms, together with the carbon atom to which they are attached. The bond distances and bond angles of the epoxide function were taken from previously reported molecular dynamics simulations of epoxides (Schiott & Bruice, 2002). The charges of the epoxide were determined from ab initio calculations using the Hartree-Fock approach, resulting in a charge of -0.46 on the oxygen atom and 0.23 on the Cα and Cβ atoms. The charges of ionizable groups of the protein were appropriate for pH 7. Arg and Lys were protonated, whereas Asp and Glu were unprotonated. Protonated forms of histidines were based on local interactions.

All simulations were performed using the GROMACS (version 3.0) package (Berendsen et al., 1995; Lindahl et al., 2001; van der Spoel et al., 2001) in a periodic dodecahedral box. Nonbonded interactions were evaluated using a twin range cutoff. Interactions within the short-range cutoff (0.9 nm) were evaluated every step, whereas interactions within the long-range cutoff (1.4 nm) were updated every five steps, together with the pair list. To correct for the neglect of electrostatic interactions beyond the 1.4 nm cutoff, a reaction field (RF) correction with εRF = 78.0 was used. To maintain a constant temperature and pressure, a Berendsen thermostat (Berendsen et al., 1984) was applied. The protein, epoxide, solvent and ions were independently coupled to a temperature bath (25 ºC) with a coupling time of 0.1 ps. The pressure was held at 1 bar, with a coupling time of 0.5 ps. The isothermal compressibility was 4.6 x 10-5 bar-1 (Berendsen et al., 1984). The time step was 0.002 ps. The bond lengths and angle in water were restrained using the SETTLE algorithm (Miyamoto & Kollman, 1992). Bond lengths within the protein were constrained using the LINCS algorithm (Hess et al., 1997).

Free energy calculations

The difference in Gibbs free energy between two states of a system was determined using the coupling parameter approach in conjunction with the thermodynamic integration (TI) formula (Mark, 1998):

λA δH (λ) ∆GA→B = λ dλ (1) ∫λB δλ

In this approach, the Hamiltonian H is made a function of a coupling parameter, λ. The λ-dependence of the Hamiltonian defines a pathway, which connects two states A and B of the system. To solve eq 1, the ensemble average at a number of discrete λ- points was obtained by performing separate simulations for each chosen λ-point, and the integral was determined numerically. The change in Gibbs energy resulting from the change of (R)-pNSO (state A) to (S)-pNSO (state B) was achieved by gradually mutating the epoxide oxygen atom of (R)- pNSO into a carbon atom, and the Cβ atom into an oxygen atom. The interchange of the epoxide oxygen and Cβ atom thus provides a minimal way to invert the stereo center of the epoxide Cα atom. The masses of the atoms were not changed. The non-bonded

80 interactions between state A and B were interpolated using a soft-core potential (Beutler et al., 1994), as implemented in the GROMACS simulation package (Lindahl et al., 2001; Villa & Mark, 2002). Prior to the thermodynamic integration calculations, a 2 ns simulation of the

HheC:H2O:(R)-pNSO complex was performed to equilibrate the structure. The structure of the enzyme retains its overall conformation during the simulation, having an average r.m.s.d. of 0.015 nm for all the backbone atoms with respect to the crystal structure. Furthermore, the bound (R)-pNSO molecule retains its hydrogen bonds with the catalytic residues Ser132 and Tyr145. The TI calculations were subsequently started from the equilibrated structure. Separate simulations were performed at 21 discrete λ-points from λ = 0 (state A) to λ = 1 (state B). At each λ-point, the data for the complex were collected for

180 ps. To obtain ∆GA-B, the average 〈δH(λ)/δλ〉λ, at each λ-point (eq 1) was integrated using the trapezoidal method. The error in 〈δH(λ)/δλ〉λ was estimated using a block averaging procedure (Bishop & Frinks, 1987; Allen & Tildesley, 1987), and the errors were integrated to give the total error in ∆GA-B.

Results and discussion

Structure of an HheC:H2O:(R)-pNSO complex at 1.7 Å resolution

Whereas cocrystallizing HheC together with Cl- and (R)-styrene oxide (SO) previously resulted in a HheC:Cl-:(R)-SO complex (de Jong et al., 2003), cocrystallization in presence of the favored epoxide (R)-pNSO and cyanate (OCN-) resulted in a

HheC:H2O:(R)-pNSO complex, with a water molecule occupying the halide-binding site (Figure 1C). Initially, these experiments aimed to investigate the structural details of the inhibition of the dehalogenation reaction by cyanate ions (I50 = 4.5 mM) (Lutje Spelberg et al., 2002), but the cyanate ions were not found in the electron density maps. The epoxide oxygen atom of the (R)-pNSO molecule bound in the active site makes hydrogen bonds of approximately 2.9 Å with the oxygen atoms of the catalytic residues Ser132 and Tyr145. The water molecule in the halide-binding site is positioned 3.0 – 3.1 Å from the Cβ atom of the epoxide ring of (R)-pNSO. These results indicate that (R)-pNSO can also bind to the active site residues in the absence of a halide ion in the halide-binding site. This is in agreement with the capability of HheC to not only to catalyze a dehalogenation reaction, but also the reverse reaction, the epoxide ring opening of epoxides by various anionic nucleophiles (Lutje Spelberg et al., 2002). Moreover, our

81 result could suggest that epoxides can bind prior to the entering of the nucleophile in epoxide ring opening reactions, which is the reverse of the order of release in the ordered Uni Bi mechanism of the dehalogenation reaction catalyzed by HheC (Tang et al., 2003).

Structure of an HheC:H2O:(S)-SO complex

An extensive kinetic analysis of the enantioselective dehalogenation of pNSHH by HheC showed that the unfavored (S)-enantiomer has a low binding affinity for the - active site and a three times lower dehalogenation rate (KM = 430 ± 50 µM, kcat = 7 ± 1 s 1 -1 ), compared to the (R)-enantiomer (KM = 9 ± 3 µM, kcat = 22 ± 1 s ) (Tang et al., 2003). The enantioselectivity of the pseudo-reverse reaction catalyzed by the enzyme, epoxide ring opening of pNSO by azide, is even higher than for the dehalogenation reaction. The enzyme converts only (R)-pNSO to the β-azidoalcohol, while (S)-pNSO is not converted at all by the enzyme. (S)-pNSO reacts only very slowly in a non-enzymatic ring opening reaction with azide or water, yielding an α-azidoalcohol or diol (Lutje Spelberg et al., 2001).

Figure 2 (right). A) A surface representation of a part of the active site cavity of HheC with bound dehalogenation products (R)-SO and a chloride ion (de Jong et al., 2003). The oxygen atom of the epoxide ring of (R)-SO is hydrogen bonded to the side chain oxygen atoms of the catalytic residues Ser132 and Tyr145 in the upper part of the cavity. B) A surface representation of a part of the active site cavity of HheC with a bound (S)-SO molecule and a water molecule occupying the halide-binding site. The oxygen atom of the epoxide ring is hydrogen bonded to the water molecule, whereas the Cβ atom makes van der Waals interactions with the side chain oxygen atoms of Ser132 and Tyr145. The orientations of the aromatic side chains are arbitrary. The corresponding electron density is not well- defined in the crystallized complexes, suggesting that the side chain can adopt different orientations. C) Detailed view of one of the active sites in the HheC:H2O:(R)-pNSO complex with a dark colored σA-weighted 2Fo-Fc electron density map at 1.7 Å covering the water molecule and (R)-pNSO molecule contoured at 1.2 σ. The light colored mesh represents the σA-weighted Fo-Fc electron density map contoured at 3.0 σ , which was obtained after refining the structure with the unfavored (S)-pNSO. The refined distances between the different atoms in are shown in Å. D) Detailed view of one of the active sites in the HheC:H2O:(S)-pNSO complex with a dark colored σA-weighted 2Fo-Fc electron density map at 1.9 Å covering the water molecule and (R)-pNSO molecule, and contoured at 1.2 σ.

To investigate the molecular basis of the enantioselectivity of HheC towards aromatic epoxides in the ring opening reaction by azide, we tried to cocrystallize HheC with the unfavored (S)-enantiomers of SO and pNSO. In our first attempts with (S)-SO, sodium nitrite was added to the protein solution, prior to the setup of the crystallization experiments. Nitrite ions are nucleophiles of ambidental nature, which can produce different products in the enzymatic epoxide ring opening reaction, depending on the attack by either the nitrogen or oxygen atom (Lutje Spelberg et al., 2002). Attack by the nitrogen atom would yield a stable β-nitroalcohol, whereas attack by the oxygen atom would form an instable nitro-ester that is easily chemically hydrolyzed to a diol (Lutje Spelberg et al., 2002). The latter reaction would recycle the nitrite ion for a second round of catalysis. Therefore, addition of a catalytic amount of sodium nitrite can convert traces of (R)- epoxides in the almost enantiopure (S)-SO sample. In this way, the (S)-enantiomer of SO that remains in solution may be observed to be bound to HheC due to the lack of competition from the (R)-enantiomer.

Using this strategy, tetragonal crystals of a HheC:H2O:(S)-SO complex in space group P43212 were obtained that diffracted to 2.7 Å resolution. The refinement of the complex resulted in subtle differences compared to a previously characterized complex of HheC with (R)-SO (de Jong et al., 2003). In the latter HheC:Cl-:(R)-SO complex, the oxygen atom of the epoxide ring makes hydrogen bonds of 2.7 Å with the side chain oxygen atoms of Ser132 and Tyr145, whereas the Cβ atom of the epoxide group is positioned 3.1 Å from the chloride ion in the halide-binding site (Figure 2A) (de Jong et al.,

2003). In the present HheC:H2O:(S)-SO complex, the aromatic ring and epoxide group of (S)-SO are located at the same location as in the HheC:Cl-:(R)-SO complex. Refinement of this complex showed that it was incompatible with a bound (R)-SO molecule, since the (R)-SO epoxide oxygen atom moved away from the Ser132 and Tyr145 side chain oxygen atoms to significantly longer distances of 3.1 and 3.4 Å, while the epoxide Cβ atom approached a water molecule in the halide-binding site at 2.7 Å. This suggested that the electron density corresponded to a bound (S)-SO, which has the same shape as its mirror image, but with the crucial difference that the oxygen and Cβ atom of the epoxide ring are interchanged (Figure 2A & B). In this binding mode, the epoxide oxygen atom of (S)-SO is hydrogen bonded to the water molecule in the halide-binding site, whereas the Cβ atom makes van der Waals interactions with the hydroxyl groups of the catalytic serine and tyrosine residues.

Structure of an HheC:H2O:(S)-pNSO complex at 1.9 Å resolution

The limited resolution of the HheC:H2O:(S)-SO complex described above may not give sufficient evidence for the proposed binding mode of (S)-SO. However, we obtained crystals that diffracted to high resolution (1.9 Å resolution) from cocrystallization experiments of HheC in the presence of highly enantiopure (S)-pNSO (97.5% e.e.), but without the addition of sodium nitrite. From the obtained electron density maps it was immediately clear that a water molecule occupies the halide-binding site of HheC in all sixteen molecules in the asymmetric unit, and that electron density for a pNSO molecule is present in all molecules, although it is not well defined in all of them. To obtain an unbiased view of the distances of the epoxide moiety to the Ser132 and Tyr145 side chain oxygen atoms and the water molecule in the halide-binding site, we modelled the substrate as para-nitrophenyl-cyclopropane. Refinement of the HheC structure with the bound p-nitrophenyl-cyclopropane against the diffraction data of both the HheC:H2O:(S)-pNSO and HheC:H2O:(R)-pNSO complexes resulted in a clear discrimination between the two. In the HheC:H2O:(R)-pNSO complex, the cyclopropane ring has moved towards the catalytic residues, compatible with a hydrogen bonding interaction of (R)-pNSO with the catalytic residues. In contrast, refinement against the

HheC:H2O:(S)-pNSO data, the cyclopropane ring is closer to the water molecule in the halide-binding site (Figure 2C, D), as expected for a hydrogen bonded (S)-pNSO. This result was confirmed by the refinement of (R)- and (S)-pNSO against the diffraction data of the HheC:H2O:(R)-pNSO and the HheC:H2O:(S)-pNSO complexes, respectively. From this we conclude that (S)-pNSO is bound in a non-productive binding mode in the active site of HheC.

Calculations of the relative Gibbs binding energy between (R)- and (S)- pNSO

To further substantiate the non-productive binding of the (S)-enantiomers of pNSO and SO, we determined the relative Gibbs binding energy (∆∆GA→B) of the two enantiomers of pNSO from thermodynamic integration calculations. ∆∆GA→B between the HheC-bound states of (R)- and (S)-pNSO, with a water molecule in the halide-binding site, was evaluated using the thermodynamic cycle shown in Figure 3A. The calculations were simplified by the notion that mutating (R)-pNSO into (S)-pNSO in water requires zero work, showing that ∆∆G(R)→(S) is equal to ∆G(R)→(S). Therefore, we can limit our calculations to the work done to (non-physically) mutate (R)-pNSO into (S)-pNSO while

85 bound to HheC, which is accomplished by the stepwise transformation of the Cβ and O atoms of the epoxide ring into each other. The relative Gibbs binding energy is estimated from the change in potential energy that is obtained from 180 ps molecular dynamics simulations at 21 different λ-points along the pathway (Figure 3B). The numerical integration of the area underneath the curve resulted in a relative Gibbs binding energy of +4.7 ± 3.1 kJ/mol (+1.1 ± 0.7 kcal/mol) when mutating (R)- into (S)-pNSO in the complex, which is in the range of the average Gibbs energy loss upon rupture of a single peptide hydrogen bond in water (Fersht et al., 1985; Shue et al., 2003). The small difference in Gibbs binding energy is in good agreement with the ability of HheC to accommodate both enantiomers in the active site. During the simulations, the positions of the aromatic side chain remained the same, fixing the location of the epoxide ring. The epoxide rings of both enantiomers of pNSO fit snugly in the active site. Therefore, the difference in Gibbs binding energy is unlikely to depend on changed van der Waals interactions, but rather on the reduced number of hydrogen bonds that (S)- pNSO is able to make. In the (R)-pNSO complex, the oxygen function of the epoxide group is hydrogen bonded to the catalytic residues, whereas in the (S)-pNSO complex, the oxygen atom of (S)-pNSO is at hydrogen bonding distance from the water molecule in the halide-binding site. Therefore, during the simulations, (R)-pNSO retains a productive binding mode in the active site of HheC, while (S)-pNSO does not. From the relative Gibbs binding energy of the (R)- and (S)-pNSO bound states, we can in principle estimate the relative binding constants for the two enantiomers, but the large error in the outcome does not allow an accurate estimation of the relative binding constants. Further computational studies are necessary to make a more accurate estimation of the difference in the Gibbs binding energy. Currently, we are assessing the reliability of our computational approach by simulating the reverse pathway, the mutation of (S)-pNSO into (R)-pNSO in the complex, which is expected to be of equal size, but with opposite sign.

Figure 3. A) A schematic representation of the thermodynamic cycle that was used in the TI calculations. The boxed part is the only part that needs to be evaluated, because mutating (R)-pNSO in (S)-pNSO in water requires zero work. Therefore ∆∆G(R)→(S) is equal to ∆G(R)→(S). B) This plot shows the calculated average 〈δH(λ)/δλ〉λ and its error at the 21 chosen λ-points. The area underneath the curve was integrated, yielding a ∆G(R)→(S) of +4.7 ± 3.1 kJ/mol.

Conclusions

Here we have presented a combined crystallographic and computational analysis of the binding of (R)- and (S)-enantiomers of the aromatic epoxides SO and pNSO to the haloalcohol dehalogenase HheC, with a water molecule rather than a halide ion occupying the halide-binding site. The crystallographic data show that both enantiomers bind with their aromatic side chains in an identical conformation in the active site, but with the crucial difference that the oxygen and Cβ atom of the epoxide ring have taken each other's positions. The oxygen atom of the epoxide ring of the bound (S)-enantiomer is in the wrong position to accept a proton from the catalytic residue Tyr145, and its Cβ atom is too far away for attack by an azide nucleophile. The wrong orientation of the epoxide ring thus prevents the enzymatic azidolysis reaction of (S)-SO and (S)-pNSO. Calculation of the relative Gibbs binding energy of the (R)-pNSO and (S)-pNSO bound states suggests that the difference in Gibbs energy of binding between the two is relatively small (∆G(R)→(S) = + 4.7 ± 3.1 kJ/mol). On the basis of these results, we propose that the enantioselectivity of the nucleophilic ring opening reaction of aromatic epoxides by azide depends only partly on the the lower affinity of HheC for (S)-enantiomers, but also for a great extend on the low kcat due to their inability to obtain a reactive orientation in the active site.

References

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Chapter 6: The X-ray structure of trans-3- chloroacrylic acid dehalogenase reveals a novel hydration mechanism in the tautomerase superfamily

René M. de Jong, Wim Brugman, Gerrit J. Poelarends, Chris P. Whitman & Bauke W. Dijkstra

Abstract

Isomer-specific 3-chloroacrylic acid dehalogenases function in the bacterial degradation of 1,3- dichloropropene, a compound used in agriculture to kill plant-parasitic nematodes. Crystal structures of the native heterohexameric trans-3-chloroacrylic acid dehalogenase (CaaD) from Pseudomonas pavonaceae 170 and of the enzyme inactivated by 3-bromopropiolate show that Glu-52 in the α- subunit is positioned to function as the water-activating base for the addition of a hydroxyl group to C- 3 of 3-chloroacrylate and 3-bromopropiolate while the nearby Pro-1 in the β-subunit is positioned to provide a proton to C-2. Two arginine residues, αArg-8 and αArg-11, interact with the C-1 carboxylate groups, thereby polarizing the α,β-unsaturated acids. The reaction with 3-chloroacrylate results in the production of an unstable halohydrin, 3-chloro-3-hydroxypropanoate, which decomposes into the products malonate semialdehyde and HCl. In the inactivation mechanism, however, malonyl bromide is produced, which irreversibly alkylates the βPro-1. CaaD is related to 4-oxalocrotonate tautomerase (4-OT), with which it shares an N-terminal proline. However, in 4-OT, Pro-1 functions as a base participating in proton transfer within a hydrophobic active site, whereas, in CaaD, the acidic proline is stabilized in a hydrophilic active site. The altered active site environment of CaaD thus facilitates a previously unknown reaction in the tautomerase superfamily, the hydration of the α,β-unsaturated bonds of trans-3-chloroacrylate and 3- bromopropiolate. The mechanism for these hydration reactions represents a novel catalytic strategy that results in carbon-halogen bond cleavage.

Published in: Journal of Biological Chemistry, in press.

Introduction

Dehalogenases are enzymes that cleave carbon-halogen bonds. They are found in various bacteria, allowing them to use halogenated hydrocarbons as growth substrates (Janssen et al., 2001, 2003). Detailed three-dimensional structural information is available for dehalogenases such as haloalkane dehalogenase, 2-haloacid dehalogenase, and haloalcohol dehalogenase, which catalyze the cofactor-independent cleavage of the covalent bond between a halogen and an sp3-hybridized carbon atom by substitution mechanisms (Verschueren et al., 1993, Ridder et al., 1997, De Jong et al., 2003). In addition, several cofactor-dependent dehalogenases have been characterized that cleave the bond between a halogen and an sp2-hybridized carbon atom. Examples include heme- dependent reductive dehalogenases (Neumann et al., 2002, Kiefer Jr et al., 2002) and the 4-chlorobenzoyl-CoA dehalogenases (Benning et al., 1996). In contrast, cofactor- independent dehalogenases that cleave the bond between a halogen and an sp2- hybridized carbon atom have only recently been discovered (van Hylckama Vlieg et al., 1992, Poelarends et al., 1998). The 3-chloroacrylic acid dehalogenases from Pseudomonas pavonaceae 170 represent cofactor-independent dehalogenases that catalyze the cleavage of vinylic carbon-halogen bonds, in which the halogen is bound to an sp2-hybridized carbon atom (van Hylckama Vlieg & Janssen, 1992; Poelarends et al., 1998; Poelarends et al., 2001). They are part of a multi-enzyme degradation route for the cis- and trans-isomers of 1,3- dichloropropene (DCP). Cis- and trans-DCP are components of the commercially produced fumigants Shell D-D and Telone II, which are used in agriculture to control plant- parasitic nematodes (Poelarends et al., 1998). The first step in the degradation of both DCP isomers is the hydrolytic cleavage of the bond between the chlorine and the sp3- hybridized carbon atom by a haloalkane dehalogenase (Figure 1). Oxidation of the two products yields the cis- and trans-isomers of 3-chloroacrylic acid, which are subsequently dehalogenated by either the cis- or trans-specific 3-chloroacrylic acid dehalogenase, yielding malonate semialdehyde and HCl. After enzymatic decarboxylation of malonate semialdehyde by malonate semialdehyde decarboxylase (Poelarends et al., submitted), the product, acetaldehyde, is likely channeled into the Krebs cycle. The trans-specific 3-chloroacrylic acid dehalogenase (CaaD) from P. pavonaceae 170 consists of an α and a β chain having 75 and 70 residues, respectively, which both share about 25% sequence identity with the 4-oxalocrotonate tautomerase (4-

OT) from Pseudomonas putida mt-2 (Poelarends et al., 2001; Chen et al., 1992; Whitman, 2002). 4-OT is a trimer of homodimers of two identical 62 amino acid chains, containing six equivalent active sites (Subramanya et al., 1996). Like other structurally related members of the tautomerase superfamily (Murzin, 1996; Almrud et al., 2002), the hydrophobic active site of 4-OT harbors a catalytic N-terminal proline residue (Pro-1) with an unusually low pKa of ~6.4 that functions as a proton-transferring base in a tautomerization reaction (Stivers et al., 1996a &b; Czerwinski et al., 1997, 1999, 2001; Harris et al., 1999).

Figure 1. The degradation of trans-1,3-dichloropropene in P. pavonaceae 170. Two dehydrogenases catalyze the oxidation of the secondary alcohol, which is formed as the result of the first dehalogenation step catalyzed by a haloalkane dehalogenase. The trans-3-chloroacrylic acid dehalogenase subsequently catalyzes the second dehalogenation step, after which decarboxylation yields the product acetaldehyde (Poelarends et al., 1998).

Biochemical characterization of CaaD uncovered a new reaction specificity in the tautomerase superfamily: the dehalogenation of chloroacrylic acids (Poelarends et al., 2001). Although both the α and β chain of CaaD have an N-terminal proline, only βPro-1 is essential for catalysis (Poelarends et al., 2001). Therefore, analogous to the oligomeric structure of 4-OT (Subramanya et al., 1996), CaaD was postulated to be a trimer of αβ dimers having three functional active sites, with βPro-1 functioning as a water-activating catalytic base (Poelarends et al., 2001). Here, we present the crystal structures of native CaaD and of the enzyme inactivated by 3-bromopropiolate at 2.6 Å and 2.3 Å resolution, respectively, suggest that a glutamate, together with the N-terminal proline, catalyzes the addition of water to the α,β-unsaturated bonds of trans-3-chloroacrylate and 3-bromopropiolate via a Michael addition. Hydration of trans-3-chloroacrylate results in the formation of an unstable halohydrin intermediate, decomposition of which is accompanied by carbon-halogen bond

93 cleavage. Dehalogenation by hydration represents a novel strategy for the cleavage of vinylic carbon-halogen bonds.

Methods

The CaaD mutants αR8A, αE52Q, and αE52D were constructed using the coding sequence for the dehalogenase in plasmid pET44T2 (Poelarends et al., 2001) as the template. The αR8A mutant was generated by PCR using the primer 5′- CACGGCATATGCCGATGATCTCTTGCGACATGGCCTATGGGAGAACA-3′, where the NdeI site is in bold and the mutated codon is underlined. This primer corresponds to the 5′ end of the wild-type coding sequence and was used in combination with primer R: 5′- TTGCCCAAGCAGAGGGATCCCCTAGCT-3′, where the BamHI site is in bold. The αE52Q and αE52D mutants were generated by overlap extension PCR (Ho et al., 1989). Primer F: 5′-CACGGCATATGCCGATGATCTCTTGCGAC-3′, where the NdeI site is in bold, and primer R were used as the external primers. For the αE52Q mutant, the internal PCR primers were oligonucleotides 5′- AGCGGGATCAACTTCGTTCAGCACGGCGAGCAT-3′, where the mutated codon is in bold, and 5′-AACGAAGTTGATCCCGCT-3′ (primer A). For the αE52D mutant, the internal PCR primers were oligonucleotide 5′-AGCGGGATCAAC TTCGTTGACCACGGCGAGCAT-3′, where the mutated codon is in bold, and primer A. The PCR reactions were carried out as described before (Poelarends et al., 2001), the products purified and cloned into plasmid pET3a (Promega Corp., Madison, WI) for overexpression of the mutant genes. The mutant genes were sequenced in order to verify that only the intended changes had been introduced. Expression and purification of CaaD and its three mutants were done according to literature procedures (Poelarends et al., 2001; Wang et al., 2003). The purified mutant proteins were analyzed by electrospray ionization mass spectrometry (ESI-MS) and gel filtration chromatography as described before (Wang et al., 2003). The activity assays with the purified enzymes were performed at 22°C by following the decrease in absorbance at 224 nm, which corresponds to the hydration of trans-3-chloro- (ε = 4900 M-1 cm-1) and trans-3-bromoacrylate (ε = 9700 M-1 cm-1) (25), or by following the release of halide using a colorimetric assay (Poelarends et al., 2001). Crystals of native CaaD were obtained from 2 µL hanging drops consisting of equal amounts of protein solution (8 mg/mL) and well solution containing 22% (w/v)

PEG4000 as precipitant, 100 mM sodium acetate buffer, pH 4.8, 0.15 M ammonium acetate and 50 mM KCl as an additive. Covalently inactivated CaaD was obtained after addition of a 25-fold excess of 3-bromopropiolate to the protein solution, which was first diluted to 0.4 mg/mL to prevent overheating of the sample. The inhibitor was synthesized by a literature procedure (Strauss et al., 1930). Crystals of inactivated CaaD were obtained using the same crystallization conditions, but without added KCl. In both cases, plates of dimensions 1 x 2 x 0.2 mm3 grew in a few days. Diffraction data sets were collected in house on a MacScience image plate system using Cu-Kα radiation from a rotating anode generator. The data were processed using DENZO and SCALEPACK (Otwinowski & Minor, 1997). The native crystal diffracted to 2.6 Å resolution and belonged to space group P212121 with cell axes a = 69.8 Å, b =

100.4 Å, and c = 109.0 Å, with two (αβ)3 hexamers in the asymmetric unit. The crystal of inactivated CaaD diffracted to 2.3 Å resolution and had space group P21 with cell axes a =

55.4 Å, b = 100.6 Å, and c = 69.9 Å, and β = 98.9°. It also contained two (αβ)3 hexamers in the asymmetric unit. Molecular replacement solutions for both data sets were obtained with the program AMORE available in CCP4 (Navaza & Saludjian, 1997; CCP4, 1994). A search model was constructed from the atomic coordinates of hexameric 4-OT from P. putida mt-2 (PDB- code 1OTF, Subramanya et al., 1996), by selecting for each monomer only residues 1 to 45, which are most similar. All residues were changed to alanines except for the proline and glycine residues. Rotation and translation functions were calculated using data between 8 and 4 Å resolution. The molecular replacement yielded the position and orientation of both hexamers in the asymmetric unit. Refinement of the solutions for the inactivated enzyme by AMORE gave a correlation coefficient of 0.42 and an R-factor of 49.7%. The electron density maps at this stage were not interpretable, but some density was extending from some of the alanines of the search model. A σA-weighted map (Read, 1998) calculated from the refined solutions from AMORE was used in a combined NCS averaging, density modification and phase extension procedure using the prime-and- switch method available in the program RESOLVE (Terwilliger, 2001). This improved the overall figure of merit from 0.17 after calculation of the starting map, to 0.48 in the final cycle of the density modification procedure. The resulting σA-weighted maps from RESOLVE showed improved electron density for the amino acid side chains and allowed identification of the α and β chains in the heterodimers of CaaD. Manual building of side chains and missing main chain atoms in the electron density maps of the inactivated

95 enzyme was subsequently alternated with simulated annealing and minimization runs in CNX (Accelrys Inc., San Diego). RESOLVE was used to obtain the most unbiased maps for model building, until σA-weighted 2Fo-Fc maps were of superior quality. The final model of inactivated CaaD was used for molecular replacement using the data of the crystal of the native enzyme. The structures were built using QUANTA (Accelrys Inc., San Diego) and XtalView (McRee, 1999). Coordinates for the adduct were based on malonate and minimized in QUANTA (Accelrys Inc., San Diego) and its parameters were generated using the Hic-Up server (Kleywegt & Jones, 1998). The quality of the final models of native and inactivated CaaD was analyzed with PROCHECK (Laskowski et al., 1993). The refinement statistics and geometric quality of the models are summarized in Table 1.

Results and Discussion Crystallization and structure elucidation Although plate-like crystals of native and inactivated CaaD could easily be obtained from various acidic buffers and different PEGs as a precipitant, most of them showed smeared diffraction spots due to disorder in the plane of the crystals. Ultimately, two crystals, one of the native enzyme and one of the inactivated enzyme, were obtained that did not show these features. After molecular replacement, the structure was built from the 2.3 Å resolution electron density maps of the crystal of the inactivated enzyme, resulting in a well-defined model of the first sixty residues of α and β chains of the enzyme. The refined model of the inactivated form of CaaD was used to solve the molecular replacement using the data of the crystal of the native enzyme. Half of the diffraction images of the native crystal suffered from ice formation on the crystal, resulting in an elevated Rsym and refinement R factors. Fifteen C-terminal residues of the α chains, and ten C-terminal residues of the β chains, accounting for over 17% of the total residues, are undefined in the electron density maps of both crystals. High flexibility of these residues could provide an explanation for the disorder in the crystals.

Data statistics Native CaaD Inactivated CaaD

Space group P212121 P21

No. hexamers / ASU (chains / ASU) 2 (12) 2 (12)

Unit cell dimensions (Å, º) a = 69.8 a = 55.4 b = 100.4 b = 100.6 c = 109.0 c = 69.9 α = β = γ = 90 α = 90 β = 98.9 γ = 90

Resolution (Å) 35 – 2.7 35 - 2.3 a Rsym (%) overall (outer shell) 14.4 (39.8) 8.9 (27.7) Completeness (%) overall (outer shell) 95.6 (97.6) 98.8 (95.2) I/σ overall (outer shell) 8.5 (3.1) 11.0 (3.0) Reflections total (unique) 343897 (20766) 330560 (35250)

Refinement

Total non-hydrogen atoms 5438 5584 Total water molecules 88 212 b R/Rfree 29.2 / 35.0 22.1 / 27.4 R.m.s.d. bonds (Å) / angles (º) 0.008 / 1.32 0.006 / 1.26 R.m.s.d. B (Å2) (main chain/side chain) 0.89 / 1.34 1.23 / 2.00

Ramachandran plot (%) 93.9 97.9 Favored 5.8 2.1 Allowed 0.2 - Generously allowed 0.2 - Disallowed a Rsym = Σ|I - |/ΣI, where I is the observed intensity and the average intensity. b R = R based on 95% of the data used in refinement. Rfree = R based on 5% of the data withheld for the cross-validation test.

Table I. Data collection and refinement statistics.

Overall structure of CaaD

The crystal structure of CaaD confirms that the enzyme is a trimer of αβ heterodimers. In both crystal forms, the asymmetric unit contains two such hexamers, which superimpose with r.m.s.d.’s between 0.3 and 0.4 Å for six times 55 Cα atoms of their core structure. The dimers are formed by the antiparallel interaction of a two- stranded parallel β-sheet of one chain with the equivalent β-sheet of the other chain, forming a four-stranded β-sheet (Figure 2a). Two α-helices, one from each chain, lie antiparallel to each other at the concave side of the β-plane. Identical chains in the asymmetric unit superimpose with r.m.s.d.’s between 0.20 and 0.35 Å for 55 Cα atoms, whereas the α and β chains superimpose with r.m.s.d.'s between 1.3 and 1.5 Å. Three of these αβ heterodimers, related by a non-crystallographic three-fold rotation axis, form a barrel-like hexamer (Figure 2b). Most dimer-dimer contacts in the hexamer are mediated by the edges of the β-sheets and are mainly of a hydrophobic nature. Each β-sheet further contributes negatively and positively charged amino acids that interact with each other in the central cavity in the hexamer (Figure 2b). Additional dimer-dimer interactions are provided by a small β-hairpin structure at the C-terminus of the α chains, extending the β- sheet of a neighboring dimer, and partly covering the environment of the catalytic βPro-1. A similar β-hairpin structure is observed in the β chains, which contributes to the environment of the catalytically inactive Pro-1 of the α chain.

The active site of native CaaD

CaaD contains three active sites related by the three-fold rotation of its (αβ)3 trimeric structure. They are located at the interface between two (αβ) dimers on one side of the hexamer, each harboring the catalytically important N-terminal proline of the β chain (Figure 2b). The N-terminal proline residues of the α chains are at the opposite side of the hexamer and are also related to each other by the (αβ)3 three-fold rotation. However, the environments of the αPro-1 and βPro-1 are different. The catalytically active βPro-1 is located in the interface between two (αβ) dimers with several charged residues in its vicinity. Among them are the side chains of αArg-8 and αArg-11 from the same αβ dimer containing the catalytic βPro-1 (Figure 3a). They are situated above the βPro-1, and bind a chloride ion, possibly from the NaCl in the crystallization solution. A third charged residue near βPro-1 is αGlu-52 from the same α chain as αΑrg-8 and αArg-11 (Figure 3a). Its carboxylate group is partly disordered in the

98 native structure, but in one active site of each of the two (αβ)3 hexamers in the asymmetric unit its side chain Oε2 atom is fixed by the amino group of the amide side chain of βAsn-39. Furthermore, a water molecule bridges the glutamate and the prolyl nitrogen. The residue in the α chain equivalent to the βAsn-39 (αPhe-39) also contributes to the environment of βPro-1 by forming part of the active site wall. Unlike βPro-1, the catalytically inactive αPro-1 is mostly buried from the solvent and is surrounded by mainly apolar residues. The residues near αPro-1 that are equivalent to αΑrg-8, αArg-11 and αGlu-52, are βAla-8, βLeu-11 and βIle-52. These differences between the α and β chain result in a completely different environment for αPro-1, which accounts for the observation that the N-terminal proline of the α-subunit is not able to support the dehalogenation reaction (Poelarends et al., 2001).

Figure 2. Stereo views of (A) the dimeric and (B) the hexameric structure of CaaD. Dark colored chains in each dimer represent the β chains, whereas the light colored chains represent the α chains. The catalytically active N-terminal proline of the β chain is shown in CPK representation.

The active site of inactivated CaaD

Further insight into the catalyzed reaction was obtained from the crystal structure of an irreversibly inactivated form of CaaD, covalently modified by 3-bromopropiolate. The design of this compound was based on the known ability of fumarase to hydrate acetylene dicarboxylate (Marletta et al., 1982). The product, hydroxyfumarate, ketonizes to afford oxaloacetate. Likewise, hydration of 3-bromopropiolate produces an unstable enol, which rearranges to an acyl bromide (Marletta et al., 1982). The electron density map of the inactivated form of CaaD demonstrates the covalent modification of only βPro-1, and not αPro-1. The electron density also allowed us to unambiguously establish the structure of the covalent adduct (Figure 3b). It was first noted that none of the active sites showed electron density for a bromide ion or a bromine atom on the inhibitor. This is consistent with the previous observation of bromide release from the inhibitor upon incubation with the enzyme (Wang et al., 2003). Second, the first two atoms attached to the prolyl nitrogen resemble a carbonyl group of a peptide bond, being in one plane with the nitrogen and carbon Cα and Cδ atoms of the proline ring. From this peptide, additional electron density extends towards the side chains of αArg-8 and αArg-11 that is compatible with a carboxylate group of the adduct. The dihedral angle between the peptide plane and the plane of the carboxylate group is 67.1°, demonstrating that no unsaturated carbon-carbon bonds are present in the adduct. On the basis of these observations, we conclude that the reaction of the enzyme with 3-bromopropiolate has resulted in the attachment of a malonyl group to the nitrogen atom of the N-terminal proline (Figure 3c). This result is in agreement with the previously reported mass increase of the β-chain upon covalent modification (Wang et al., 2003). The modification of the βPro-1 by a malonyl group may at first seem surprising, but is readily explained by the addition of a water molecule to the carbon-carbon triple bond as a first step in the reaction of 3-bromopropiolate with the enzyme (see below).

Figure 3 (right). A) Stereo view of the active site of native CaaD, showing the chloride ion (green) bound to αArg-8 and αArg-11 and the phenylalanine (αPhe-50) and the glutamate (αGlu-52) of the β- hairpin of the α-chain, binding to βAsn-39 in close proximity to βPro-1. B) The final 2Fo-Fc electron density map from XtalView (31) covering the proline and the covalently bound adduct, clearly showing the planar peptide link to βPro-1 and the twist of the carboxyl plane with respect to the peptide plane. C) Active site of the inactivated enzyme showing the carboxylate group of the covalent adduct interacting with the two arginines. D) Close-up stereo view of the superposed active site regions of inactivated CaaD and 4OT. Dark colored chains in each dimer represent the β chains, whereas the light colored chains represent the α chains of CaaD.

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101

Apart from the interaction of its carboxylate group with αΑrg-8 and αArg-11, the adduct further has hydrophobic contacts with αPhe-50, αLeu-57 and βIle-37. Interestingly, one active site in each of the two different (αβ)3 hexamers in the asymmetric unit of inactivated CaaD shows a water molecule bound between the αGlu-52 Oε2 atom and the nitrogen atom of the modified proline, at a similar position as the one observed in the native structure (Figure 3a). Strikingly, the covalent adduct in this active site shows negative electron density at the position of its carboxylate group. Because β-ketoacids readily undergo decarboxylation (Jencks, 1987), the binding of the water molecule likely correlates with the presence of an acetylated adduct resulting from decarboxylation of the malonyl group. This observation is also consistent with previous mass spectral analysis showing decarboxylation of the malonyl adduct (Wang et al., 2003). In the active sites that lack a bound water molecule, the density for the carboxylate group is fully present, indicating that decarboxylation had not occurred. In these active sites, the αGlu-52 Oε2 atoms make a hydrogen bond of approximately 2.9 Å with the backbone carbonyl group of βIle-37, suggesting that the glutamate side chain is protonated.

Mutagenesis of αArg-8 and αGlu-52

On the basis of sequence analysis and site-directed mutagenesis studies, βPro-1 and αArg-11 were previously identified as critical residues in CaaD (Poelarends et al., 2001; Wang et al., 2003). The crystallographic results suggest that αArg-8 and αGlu-52 may also play important roles in the mechanism. Therefore, three single site-directed mutants were constructed in which αGlu-52 was replaced with a glutamine (αE52Q) or aspartate (αE52D) and αArg-8 with an alanine (αR8A). The three mutants were expressed in E. coli BL21(DE3) and purified to ~95% homogeneity (as assessed by sodium dodecyl sulfate polyacrylamide gel electrophoresis) using the protocol described for wild-type CaaD (Wang et al., 2003). Analysis of the mutant proteins by ESI-MS and gel filtration chromatography indicated that the α- and β-subunits had the expected masses (and were not blocked by the initiating methionine) and that the mutants were heterohexamers in solution. In a spectrophotometric assay, which measures the depletion of substrate at 224 nm, activity was not detected for any of the mutants using trans-3-chloro- and trans-3- bromoacrylate (after 30 min). However, a 24 h incubation period of the mutant proteins (in separate reactions) with trans-3-chloroacrylate showed that the αE52D and αR8A

102 mutants retained a small amount of activity (as determined by a colorimetric assay that monitored the release of the halide) while the αE52Q mutant had no detectable activity. Based on these results, we conclude that αArg-8 and αGlu-52 are indeed important for the CaaD-catalyzed hydration reactions.

Similarity to other members of the 4-OT family

The crystal structures of the irreversibly inactivated CaaD and 4-OT (Taylor et al., 1998) superimpose with an r.m.s.d. of 2.1 Å for six times 50 Cα atoms per hexamer. The α and β chains of CaaD superimpose on the single chain that builds the 4-OT homohexamer with r.m.s.d.'s of approximately 1.3 Å and 1.6 Å, respectively, for 55 Cα atoms. The high r.m.s.d.'s are mainly due to deviations in the regions that contribute to the active site environment of βPro-1, whereas most of the β-α-β structural core of the αβ dimers superimposes nearly perfectly. In particular, the C-terminal β-hairpin structures and the loop regions between residues 35 and 41 in both chains are shifted in the direction of βPro-1 with respect to their equivalents in 4-OT, resulting in a more narrow active site cavity in CaaD. This is not surprising, since trans-3-chloroacrylate is considerably smaller than the elongated 2-oxo-4-trans-hexenedioate substrate of 4-OT. Comparison of the inhibited CaaD and 4-OT structures (Figure 3d) shows that the inhibitors interact differently with the enzymes. In 4-OT, Arg-39 binds the keto and carboxylate group of the inhibitor, while in CaaD Arg-39 is replaced by αPhe-39 and βAsn-39, and αArg-8 and αArg-11 interact with the inhibitor’s carboxylate group. Arg-11 is also present in 4-OT, where it likely interacts with the C-6 carboxylate group of 2-oxo-4- trans-hexenedioate (Harris et al., 1999). Thus, of the two arginines that bind the substrate in 4-OT, only αArg-11 is conserved in CaaD. The most striking difference between CaaD and 4-OT is the presence of αGlu-52 at 4.5 – 6.0 Å from βPro-1. In 4-OT an isoleucine is present at the equivalent position, 8 – 9 Å away from the N-terminal proline. In addition, Phe-50 contributes to the hydrophobic active site environment of Pro-1 in 4-OT; αPhe-50 is also present in CaaD. In both enzymes, these residues are part of the C-terminal β-hairpin structure that covers the active site pocket. Whereas the hydrophobic residues of the β-hairpin are important for the low pKa of the Pro-1 of 4-OT by preventing water from entering the active site (Czerwinski et al., 2001), the corresponding β-hairpin glutamate in CaaD creates a more hydrophilic active site environment. A negatively charged residue near the amino-terminal proline is

103 unprecedented in the tautomerase superfamily (Subramanya et al., 1996; Taylor et al., 1999; Whitman, 2002), and has important consequences for the role of βPro-1 in the catalytic mechanism of CaaD as well as for the mechanism of inactivation of CaaD by 3- bromopropiolate.

Mechanism of inactivation of CaaD by 3-bromopropiolate

CaaD is well equipped to catalyze the addition of water to the triple bond of 3- halopropiolates, which are analogues of the substrates for CaaD. The specific interaction of the covalent malonyl adduct with the two arginines suggests the favored orientation of 3-bromopropiolate in the active site. The bromine atom may bind deep in the active site pocket, possibly interacting with the two phenylalanines that constitute the active site wall. This would position the C3 carbon atom of the triple bond close to the water molecule that bridges αGlu-52 and βPro-1 in the native structure of CaaD, whereas the C2 carbon atom is close to the prolyl nitrogen of βPro-1. The glutamate side chain interacts with the amino group of the amide side chain of αAsn39, the bridging water molecule and a second surface-bound water molecule, suggesting that its side chain carboxylate group is likely stabilized in a charged anionic form. The water-mediated interaction of the charged glutamate with the prolyl nitrogen of βPro-1 suggests that the proline is protonated at physiological pH, which is consistent with its position in a hydrophilic environment. In contrast to the Pro-1 catalytic base in 4-OT (Czerwinski et al., 2001), βPro-1 may thus function as a general acid that protonates the C2 carbon atom of the inhibitor (Figure 4a). The anionic αGlu-52 could act as the general base that activates a water molecule for a nucleophilic attack at the C-3 position. The arginine residues, αArg-8 and αArg-11, favor the formation of a partial positive charge at C-3 by polarizing the carboxylate group of the inhibitor, thereby increasing the susceptibility of the C-3 position for nucleophilic attack. Strikingly, the glutamate is hydrogen bonded to the backbone carbonyl function of βIle-37 in the majority of the active sites of the inactivated enzyme, whereas similar interactions with hydrogen bond acceptors are absent in the native structure. This suggests that the side chain carboxylate group has been protonated during the inactivation reaction, further supporting its role as the water-activating base.

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Figure 4. A) Schematic representation of the proposed mechanism of inactivation of CaaD by 3- bromopropiolate, which results from the formation of an reactive acyl bromide intermediate that is produced by hydration of the substrate analogue. B) Schematic representation of the proposed dehalogenation mechanism of trans-3-chloroacrylate by CaaD, in which hydration of the substrate yields an instable halohydrin that subsequently collapses to yield malonate semialdehyde and HCl.

Hydration of the triple bond in 3-bromopropiolate would thus yield 3-bromo-3- hydroxypropenoate, which rapidly rearranges to yield malonyl bromide (Figure 4a). The acyl bromide function is very reactive towards nucleophiles (Bruice, 2003). Because the prolyl nitrogen of βPro-1 has lost its proton in the hydration step, it may now function as a nucleophile and react with the acyl bromide, resulting in the formation of the covalent malonyl adduct and the departure of a bromide ion (Wang et al., 2003). It was previously proposed that inactivation of CaaD by 3-bromopropiolate could occur by this mechanism- based route to produce an acyl bromide (or a ketene). Alternatively, a Michael addition of βPro-1 to C-3 of 3-bromopropiolate was proposed (Wang et al., 2003). The crystallographic results strongly support the mechanism-based route, because the active

105 site is clearly set up to carry out a hydration reaction and βPro-1 is not sufficiently nucleophilic to attack 3-bromopropiolate in a Michael fashion. Thus, 3-bromopropiolate is a mechanism-based inhibitor of CaaD.

Conversion of trans-3-chloroacrylate by CaaD

The geometry of the inactivated complex, the mechanism proposed for inactivation, and the results of the site-directed mutagenesis experiments suggest a mechanism for the dehalogenation of trans-3-chloroacrylate by CaaD. In the catalytic mechanism, αGlu-52 and βPro-1 may catalyze the Michael addition of water to the double bond of trans-3-chloroacrylate (Figure 4b). The two arginines likely polarize the carboxylate group, thereby facilitating nucleophilic attack at C-3. The addition of water results in the unstable 3-chloro-3-hydroxypropanoate intermediate, which subsequently decomposes to yield malonate semialdehyde and HCl. The cleavage of the carbon- chlorine bond in the chlorohydrin intermediate is now a chemically favorable reaction that may not need to be further catalyzed by specific halide-binding residues (Marletta et al., 1982). There is, however, a possibility that the N-terminal proline residue plays an active role in the collapse of 3-chloro-3-hydroxypropanoate by deprotonating its hydroxyl group. In this manner, the proline would also regenerate its initially charged form, thereby returning the system to its native state to enable another turnover. Thus, dehalogenation of trans-3-chloroacrylate by CaaD is mechanistically a hydration reaction, which represents a novel strategy for the cleavage of a bond between a halogen and an sp2- hybridized carbon atom. An alternate mechanism involves the formation of a carbocation intermediate. In this scenario, βPro-1 initiates the reaction by providing a proton to C-2, which generates a positive charge at C-3. The nearby water molecule, held in position by αGlu-52, adds to the carbocation to afford the halohydrin intermediate. In this mechanistic scenario, the two arginines are more likely to play a role in substrate alignment. While carbocation intermediates are rare in enzyme-catalyzed reactions, such a possibility cannot be ruled out.

106

The evolutionary origin of chloroacrylate dehalogenating activity

While other structurally characterized members of the tautomerase superfamily contain homo- or fused heterodimers in different stoichiometries (Murzin, 1996; Subramanya et al., 1996; Almrud et al., 2002; Whitman, 2002), CaaD is built up of a trimer of αβ dimers of two different tightly packed peptide chains. The α and β chains form three identical active sites that carry out the dehalogenation of trans-3-chloroacrylate by hydration. Whereas most superfamily members known so far use the conserved N- terminal proline as a proton-transferring catalytic base in tautomerization reactions, the βPro-1 of CaaD functions as a general acid in a hydration reaction. Surprisingly, 4-OT and YwhB, a 4-OT homologue in Bacillus subtilis, have a rudimentary trans-3-chloroacrylic acid dehalogenase activity, in addition to their major tautomerase activities (Whitman, 2002). Such catalytic promiscuity has also been observed in other enzyme superfamilies, suggesting that these secondary activities could play a role in the evolution of new enzymatic activities (O'Brien & Herschlag, 1998). The low-level dehalogenase activity of 4-OT could result from the interaction of Arg11 with the carboxylate group to create a partial positive charge at C-3, thereby facilitating a Michael addition of water to trans-3-chloroacrylate. The catalytic proline could assist in catalysis by activating the water molecule. The efficiency of the dehalogenation reaction might have been limited by the enzyme’s inability to carry out a hydration reaction in a hydrophobic pocket. Moreover, the presence of water in the active site could raise the pKa of the proline, thereby decreasing its effectiveness as a base. Evolutionary selection may ultimately have led to the altered active site environment that enabled a more efficient hydration of the substrate. A BLAST search (Altschul et al., 1997) against the protein sequence databases yields various (hypothetical) 4-OT sequences. Only one of them shares the characteristic glutamate and two arginines that are present in CaaD, suggesting that this variation in the tautomerase superfamily is rare. The gene (cg10062), encoding a 149 amino acid protein corresponding to a fused α and β chain of CaaD, occurs in the genome of Corynebacterium glutamicum ATCC 13032 (Ikeda & Nakagawa, 2003), a bacterium that is widely used in the amino acid fermentation industry. Interestingly, this gene product exhibits a low-level dehalogenase activity towards cis-3-chloroacrylate (Poelarends GJ, Serrano H and Whitman CP, unpublished results). This may be another example of a hydratase or tautomerase with promiscuous dehalogenase activity.

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Poelarends GJ, Saunier R, Janssen DB: trans-3-Chloroacrylic acid dehalogenase from Pseudomonas pavonaceae 170 shares structural and mechanistic similarities with 4- oxalocrotonate tautomerase. J Bacteriol 2001, 183:4269-4277. Poelarends GJ, Wilkens M, Larkin MJ, van Elsas JD, Janssen DB: Degradation of 1,3- dichloropropene by Pseudomonas cichorii 170. Appl Env Microbiol 1998, 64:2931-2936. Read RJ: Improved Fourier coefficients for maps using phases from partial structures with errors. Acta Crystallogr 1986, A42:140-149. Ridder IS, Rozeboom HJ, Kalk KH, Janssen DB, Dijkstra BW: Three-dimensional structure of L-2- haloacid dehalogenase from Xanthobacter autotrophicus GJ10 complexed with the substrate analogue formate. J Biol Chem 1997, 272:33015-33022. Stivers JT, Abeygunawardana C, Mildvan AS, Hajipour G, Whitman CP, Chen LH: Catalytic role of the amino-terminal proline in 4-oxalocrotonate tautomerase: affinity labeling and heteronuclear NMR studies. Biochemistry 1996, 35:803-813. Stivers JT, Abeygunawardana C, Mildvan AS, Hajipour G, Whitman CP: 4-Oxalocrotonate tautomerase: pH dependence of catalysis and pKa values of active site residues. Biochemistry 1996, 35:814-823. Strauss F, Kollek L, Heyn W: Über den Ersatz positiven Wasserstoffs durch halogen. Chem Ber 1930, 63:1868-1899. Subramanya HS, Roper DI, Dauter Z, Dodson EJ, Davies GJ, Wilson, KS, Wigley DB: Enzymatic ketonization of 2-hydroxymuconate: specificity and mechanism investigated by the crystal structures of two isomerases. Biochemistry 1996, 35:792-802. Taylor AB, Czerwinski RM, Johnson WH, Whitman CP, Hackert ML: Crystal structure of 4- oxalocrotonate tautomerase inactivated by 2-oxo-3-pentynoate at 2.4 A resolution: analysis and implications for the mechanism of inactivation and catalysis. Biochemistry 1998, 37:14692- 14700. Taylor AB, Johnson Jr. WH, Czerwinski RM, Li HS, Hackert ML, Whitman CP: Crystal structure of macrophage migration inhibitory factor complexed with (E)-2-fluoro-p-hydroxycinnamate at 1.8 A resolution: implications for enzymatic catalysis and inhibition. Biochemistry 1999, 38:7444-7452. Terwilliger TC: Map-likelihood phasing. Acta Crystallogr 2001, D57:1763-1775. van Hylckama Vlieg JET, Janssen DB: Bacterial degradation of 3-chloroacrylic acid and the characterization of cis- and trans-specific dehalogenases. Biodegradation 1992, 2:139-150. Verschueren KH, Seljée F, Rozeboom HJ, Kalk KH, Dijkstra BW: Crystallographic analysis of the catalytic mechanism of haloalkane dehalogenase. Nature 1993, 363:693-698. Wang SC, Person MD, Johnson Jr. WH, Whitman CP: Reactions of trans-3-chloroacrylic acid dehalogenase with acetylene substrates: consequences of and evidence for a hydration reaction. Biochemistry 2003, 42:8762-8773. Whitman CP: The 4-oxalocrotonate tautomerase family of enzymes: how nature makes new enzymes using a beta-alpha-beta structural motif. Arch Biochem Biophys 2002, 402:1-13.

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Chapter 7: Outlook

111

Introduction

Answers to questions inevitably raise new questions, and established ideas, theories and concepts will always continue to be challenged at their marshy boundaries with the unknown. Some unsolved questions also remain at the end of my Ph.D. project, although our current knowledge of the different catalytic mechanisms that are utilized by the various bacterial dehalogenating enzymes has considerably expanded. The most intriguing questions that remain are discussed below, together with suggestions on how we may answer them experimentally.

What are the possible halide release pathways in HheC?

Extensive characterization of the kinetic mechanism of the dehalogenase reaction catalyzed by haloalcohol dehalogenase HheC revealed that halide-release is the rate-limiting step, and likely occurs prior to the release of the epoxide product (Tang et al., 2003). Lack of detailed understanding of the possible pathway(s) taken by the halide ion upon release, however, hampers structure-based protein design to improve the catalytic performance of HheC. Mutations of amino acids that are positioned along the release pathway may accelerate the rate-limiting halide release step, an approach that has successfully been applied to accelerate the slow halide release from the active site of haloalkane dehalogenase from Sphingomonas paucimobilis UT26 (Chaloupkova et al., 2003). From the X-ray structure of HheC, the most feasible halide release pathway seems one via the back of the halide-binding site (Figure 1a). The structure shows that there is a void created by the relatively loose packing of the amino acids that surround Leu178, suggesting that the side chain of this residue could switch to a different rotamer conformation, and allow a halide ion or nucleophile to pass. Interestingly, activity essays showed that a Leu178Phe mutant of HheC is completely inactive. An explanation for this inactivity could be that the larger and rigid aromatic side chain of the phenylalanine is more restrained by its neighboring residues, which could prevent passage of a halide ion. However, improper folding of the halide-binding site can not be excluded as the cause for the inactivity of the mutant.

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Figure 1. A) Detailed structure of the halide-binding site of HheC. B) Detailed structure of HheC in complex with the products Cl- and (R)-styrene oxide.

Intriguingly, biochemical data on other HheC mutants may suggest an alternative entrance/release pathway. Kinetic data on Tyr187Phe and Trp249Phe mutants showed an significant improvement of the catalytic activity, indicating an acceleration of the rate- limiting halide release step (Lixia Tang, unpublished results). The side chains of these two residues and an asparagine are hydrogen-bonded to each other, and form part of the active site wall near the entrance of the substrate-binding site (Figure 1b). The rate enhancement upon destabilization of the hydrogen-bonding network as a result of the mutation thus may suggest that the release pathway is located opposite to the halide- binding site, close to the entrance of the haloalcohol or epoxide substrate. From the structure of HheC, it is, however, unclear how a halide ion could be released via this side of the enzyme without significant movements of several amino acid residues. An alternative explanation for the rate improvement could be that the mutations improve the dynamics of halide-release via the pathway at the back of the halide-binding site.

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Mutants of Leu178 may offer experimental proof that could resolve the questions on the pathway of halide release. Single turn-over experiments with such mutants may discriminate between the two suggested explanations for its observed inactivity. On the one hand, the inactivity may imply that formation of the halide and epoxide products is prevented due to a misfolded halide-binding site. Alternatively, only halide-release may be impaired by the larger and more rigid aromatic side chain of the phenylalanine in the mutant. UV/VIS spectra of the dehalogenation reaction in a single turn-over experiment can distinguish between product formation and halide-release in the wild type enzyme (Tang et al., 2003). If product formation is still observed and halide-release becomes impaired when using the Leu178 mutants, this would evidence the involvement of Leu178 in halide release. Moreover, these experiments may also exclude the presence of other halide release pathways in the wild type dehalogenase.

Can we engineer a redox activity in haloalcohol dehalogenase HheC?

Our structure-based alignments of HheC and the structurally most similar SDR enzymes convincingly show that the haloalcohol dehalogenases evolved from members of different NAD-specific subfamilies of the SDR family. A challenging idea would be to explore the possibility to engineer NAD-binding affinity in the dehalogenase that could support a redox reaction, rather than a substitution reaction. Such an attempt would at least require the removal of the voluminous amino acids that replace the characteristic Gly-rich NAD(P)H-binding loop of the short-chain dehydrogenases/reductase family in the dehalogenase. Strikingly, modeling of the Gly-rich loop in the structure of the dehalogenase suggests that this is sterically well possible, and that the loop may adopt a similar conformation as generally observed in the SDR enzymes. A similarly shaped cofactor binding groove is thus obtained in the model, which can accommodate an NADH cofactor. Moreover, the model of the mutant also contains two conserved residues (Ser180, and Asp32 in HheC), which are properly positioned to make specific hydrogen bonds with the NADH cofactor, similar to the ones observed in structures of NAD-specific SDR enzymes. The modeling experiments thus strongly encourage the idea that relatively few mutations may create a functional NADH-binding site, which could possibly facilitate a dehydrogenase or reductase activity in the dehalogenase.

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Figure 2. A close-up view in stereo of the structure of HheC in complex with Cl- and (R)-styrene oxide, in which the Gly-rich NAD(P)H-binding SDR loop and NADH have been modeled in (in the center in black). His31, Asp32, and Ser180 are properly positioned to make important hydrogen bonding interactions with an NAD+ or NADH cofactor in the modeled enzyme. Most importantly, Asp32 can bind the sugar unit of the cofactor, whereas Ser180 can interact with the cofactor head group, similar as observed in NAD-dependent SDR enzymes.

Can we experimentally evolve dehalogenating activities in enzyme superfamilies?

Although the results presented in this thesis clearly evidence evolutionary relationships between haloalcohol dehalogenases and NADH-dependent SDR enzymes, and between 3-chloroacrylic acid dehalogenases and 4-oxalocrotonate tautomerases, we can only speculate about the adaptive routes along which the dehalogenation activities have emerged in the different enzyme superfamilies. A highly challenging idea would be to use directed evolution techniques to evolve dehalogenating activities from existing enzyme superfamily members with distinctly different activities. Directed evolution is a very promising technique to obtain highly efficient enzyme catalysts for green chemistry, as well as to teach us about the origin and evolution of protein function. Properties of enzymes like substrate specificity, enantioselectivity, catalytic efficiency and thermostability can be improved using various methods available (Powell et al., 2001, Farinas et al., 2001, Tao and Cornish, 2002). Moreover, directed evolution has been

115 successfully applied to make a protein scaffold bind new small molecules with nanomolar affinities (Beste et al., 1999), an achievement that was recently also obtained by computational methods (Looger et al., 2003). In contrast, successful attempts to use directed evolution to evolve novel enzyme activities have not been reported. Although de novo enzyme-like activity can be obtained in catalytic antibodies raised against transition state mimics (Hilvert, 2000), or by computational design of a protein scaffold (Bolon and Mayo, 2001), the obtained catalytic efficiencies are still disappointingly low. Ultimately, the desire is to use directed evolution to evolve novel catalytic activities with high efficiencies in existing enzyme scaffolds, thereby realizing an effective way to obtain custom-made enzymes for use in the chemical and pharmaceutical industry. The different dehalogenase-containing enzyme superfamilies may provide useful precursor enzymes that could be used as test cases to study the evolution of dehalogenase activities. This would not only confirm the evolutionary origin of dehalogenase activities, but also provide important insights into the directed evolution of novel enzymatic activities in general.

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References

Chaloupkova R, Sykorova J, Prokop Z, Jesenská A, Monincová M, Pavlova M, Tsuda M, Nagata Y, Damborský J: Modification of activity and specificity of haloalkane dehalogenase from Sphingomonas paucimobilis UT26 by engineering of its entrance tunnel. J Biol Chem 2003, Epub ahead of print. Tang L, Lutje Spelberg JH, Fraaije MW, Janssen DB: Kinetic mechanism and enantioselectivity of halohydrin dehalogenase from Agrobacterium radiobacter. Biochemistry 2003, 42:5378-5386. Powell KA, Ramer SW, Del Cardayre SB, Stemmer WP, Tobin MB, Longchamp PF, Huisman GW: Directed Evolution and Biocatalysis. Angew Chem Int Ed 2001, 40:3948-3959. Farinas ET, Bulter T, Arnold FH: Directed enzyme evolution. Curr Opin Biotechnol 2001, 12:545- 551. Tao H, Cornish VW: (2002) Milestones in directed enzyme evolution. Curr Opin Chem Biol 2002, 6:858-864. Beste G, Schmidt FS, Stibora T, Skerra A: Small antibody-like proteins with prescribed ligand specificities derived from the lipocalin fold. Proc Natl Acad Sci USA 1999, 96:1898-1903. Looger LL, Dwyer MA, Smith JJ, Hellinga HW: Computational design of receptor and sensor proteins with novel functions. Nature 2003, 423:185-190. Hilvert D: (2000) Critical analysis of antibody catalysis. Annu Rev Biochem 2000, 69:751-793. Bolon DN, Mayo SL: Enzyme-like proteins by computational design. Proc Natl Acad Sci USA 2001, 98:14274-14279.

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Chapter 8: Summary

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Introduction

It is generally believed that evolution is at the heart of species differentation of all living organisms, a hypothesis that was first stated by Charles Darwin in his revolutionary work “The Origin of Species”. Although this hypothesis was originally based on phenotypic similarities between different species, the current knowledge of the genomes of various species shows that interspecies relationships are even more evident on the genetic level, even when species have very dissimilar phenotypes. Central to the current view of Darwinian evolution and species differentiation are: (i) mutation/recombination of the genetic code and the corresponding phenotypic changes (mutation); (ii) differential survival of altered individuals in the population (selection); and (iii) the propagation of beneficial changes in the population by (a)sexual replication (amplification). Despite this concise theoretical concept, the mechanisms that gave rise to the current species diversity are poorly understood. This is partly due to the tremendously large timescale of evolutionary changes becoming apparent in higher organisms, making it impossible to get a good idea of the intermediates along the pathway. Often, fossils are the only proof of the past existence of such intermediates ('missing links') that could support the hypothesized evolutionary path between different species, like, for example, the evolution of birds from reptiles and the evolution of Homo sapiens. It is therefore not surprising that the theory of evolution has long been in the realm of theoretical biology, suffering from the inaccessibility to empirical experiments to support hypotheses. Nowadays this situation is drastically changing. The elucidation of the complete genome sequences of different organisms and the development of modern molecular biology techniques have cleared the road to experimental evolution. Many examples of evolutionary adaptation are being studied in model organisms in the laboratory, like, for example, the adaptation of pathogenic bacteria to antibiotics, or the adaptation of the HIV virus to antiviral drugs. Such studies have made clear that microbes, which divide approximately every 10 to 30 minutes, can rapidly adapt themselves to external factors by mutation and recombination of their gene repertoire. Structure determination techniques like X-ray crystallography and nuclear magnetic resonance (NMR) have revealed the molecular basis of such evolutionary adaptations, in which mutations in a gene can cause changes in the gene-encoded product. Subsequent selection and proliferation of mutated genes thus may result in changes in activity and function of proteins inside the cell, thus providing the basis of evolution at the molecular level.

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Dehalogenation of anthropogenic halogenated hydrocarbons by bacteria

Different ways to cleave a carbon-halogen bond

Biochemical characterization of several dehalogenases from bacteria revealed a wide variety of different chemical strategies that are used in nature to facilitate cleavage of carbon-halogen bonds. Several anaerobic halorespiring bacteria contain reductive dehalogenases that enable them to use tetrachloroethene, rather than oxygen, as the final electron acceptor in the respiratory chain. Aerobic bacteria, instead, may contain mono- or dioxygenases to oxidatively dehalogenate small haloalkenes, haloalkanes and some aromatic compounds. Most of the structurally characterized dehalogenases, however, utilize cofactor-independent SN2-type substitution mechanisms that proceed via a covalently bound intermediate with a nucleophilic aspartate, which is subsequently hydrolyzed by a water molecule. Haloalkane dehalogenase, haloacid dehalogenase and 4-chlorobenzoyl-coenzyme A dehalogenase all make use of such a mechanism, but have different amino acid sequences and different three-dimensional structures. They are evolutionary unrelated to large superfamilies of enzymes with fundamentally different activities. Subtle changes in the active site of the dehalogenases make the dehalogenation activity possible, but the lack of amino acid sequence identity with other

121 members of the enzyme superfamily make it impossible to identify the likely precursor enzyme.

Molecular basis of two novel dehalogenating activities in bacteria

This thesis describes the X-ray crystallographic elucidation of the molecular structures of two novel, unrelated bacterial dehalogenases, the haloalcohol dehalogenase HheC from Agrobacterium radiobacter AD1, and a trans-3-chloroacrylic acid dehalogenase (CaaD) from Pseudomonas pavonaceae 170. Structures of the native enzymes, and of complexes of the enzymes with products, substrate analogues, or inhibitors provide conclusive insights into the mechanistic details of two novel dehalogenation strategies, which fundamentally differ from the mechanisms of previously characterized haloalkane -, haloacid -, and 4-chlorobenzoyl-Coenzyme A dehalogenases. Chapter 2 gives a detailed overview of the structures and mechanisms of the different bacterial dehalogenases, emphasizing the differences and similarities with evolutionary related enzyme families, and on the possible evolutionary origins of the dehalogenases. In chapter 3, 4 and 5, the crystallization, the structure elucidation and the catalytic mechanism of haloalcohol dehalogenase HheC are described. HheC catalyzes the dehalogenation of vicinal haloalcohols like 2,3-dichloro-1-propanol via an one-step intramolecular substitution mechanism, yielding the corresponding epoxide and HCl. The results show that HheC is structurally and mechanistically related to a widespread family of NAD(P)H-dependent redox enzymes, the short-chain dehydrogenases/reductases (SDR family). X-ray structures of HheC with bound products and a substrate analogue demonstrate that catalysis depends on a conserved Ser-Tyr-Arg catalytic triad. The SDR NAD(P)H-binding site has changed, however, into a halide-binding site that supports the cofactor-independent dehalogenation reaction. Structure-based sequence alignments further evidence that the six haloalcohol dehalogenases that are currently known have evolved from two different NAD-binding SDR enzymes, rather than from NAD(P)-binding enzymes. In chapter 6, I present X-ray structures of the trans-3-chloroacrylic acid dehalogenase CaaD from P. pavonaceae 170, both of the native enzyme and of a form that is covalently inactivated by a mechanism-based inhibitor. The results provide conclusive evidence that CaaD cleaves an sp2-hybridized carbon-halogen bond via a Michael addition of water to trans-3-chloroacrylate, a degradation product of the soil

122 fumigant 1,3-dichloropropene. Subsequent decomposition of the product, an unstable halohydrin, results in the release of malonate semialdehyde and HCl. CaaD is related to the well-studied Pseudomonas enzyme 4-oxalocrotonate tautomerase (4-OT). Whereas this tautomerase uses an N-terminal catalytic proline (Pro1) as a base to transfer protons in tautomerization reactions, the Pro1 in the dehalogenase is acidic, stabilized in a hydrophilic active site. Together with a buried glutamate base, the acidic proline facilitates a hydratase activity, rather than a tautomerase activity.

On the origin of dehalogenating activities in bacteria

Although the evolutionary origins of most dehalogenases remain unclear, the structural information on haloalcohol dehalogenase HheC and trans-3-chloroacrylic acid dehalogenase CaaD offers insights into their likely evolutionary origins. In contrast with the haloalkane dehalogenases and haloacid dehalogenases, only a few amino acid sequences of haloalcohol dehalogenases and 3-chloroacrylic acid dehalogenases are known. Their rare occurrence in soil bacteria, and their absence in other organisms, suggests that they may have evolved from related enzymes with different activities in response to elevated concentrations of their halogenated substrates in the environment. Catalytic promiscuity of a precursor enzyme may have played an important role in their emergence in bacteria. An SDR precursor of the haloalcohol dehalogenases may fortuitiously have bound a haloalcohol substrate, possibly with the halogen atom positioned in the NADH cofactor-binding site. The catalytic tyrosine may have been able to deprotonate the haloalcohol hydroxyl group, thereby catalysing the intramolecular substitution reaction. Such a rudimentary dehalogenase activity may subsequently have been optimized by evolutionary selection. A similar scenario may apply to the emergence of 3-chloroacrylic acid dehalogenase activity in bacteria. In contrast to the SDR enzymes, a few tautomerases have been experimentally shown to posses a rudimentary dehalogenase activity. It will be a scientific challenge to verify such hypotheses on the evolutionary origin of dehalogenases using directed evolution.

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Chapter 9: Nederlandse samenvatting

125

Inleiding

Het wordt algemeen aangenomen dat evolutie aan de basis ligt van de diversiteit aan levende organismen op aarde, een hypothese die voor het eerst door Darwin werd geopperd in zijn visionaire werk “The origin of species”. Hoewel deze hypothese volledig gebaseerd werd op fenotypische overeenkomsten tussen verschillende soorten, laat onze huidige kennis van de genomen van verschillende organismen zien dat de evolutionaire verwantschap nog evidenter is op het DNA-niveau, zelfs als soorten fenotypisch zeer verschillen. Centraal in de huidige theorie van Darwiniaanse evolutie staan mutatie en recombinatie van de genetische code en de daarmee gepaard gaande fenotypische veranderingen (mutatie), de verschillende overlevingskansen van veranderde individuen binnen de populatie (selectie), en de verbreiding van voordelige veranderingen in de populatie (amplificatie). Ondanks dit aannemelijke theoretische concept, worden de mechanismen die aan de basis liggen van het ontstaan van de diversiteit aan verschillende diersoorten op aarde nog immer slecht begrepen. De tijdschaal waarop zulke evolutionaire processen plaatsvinden maakt het onmogelijk een beeld te vormen van de tussenstadia die tijdens het ontstaan van de ene soort uit de ander moeten hebben bestaan. Vaak vormen zeldzame fossielen het enige houvast om evolutionaire theorieën te staven, zoals in het geval van de oorsprong van de mens, en de evolutie van vogels uit reptielen. Het is daarom ook niet verrassend dat evolutieleer lang tot het gebied van de theoretische biologie behoorde, en voor een groot deel werd belemmerd door de ontoegankelijkheid van empirische experimenten om haar theorieën te ondersteunen. Tegenwoordig is dat anders. De opheldering van complete genomen van uiteenlopende organismen en de komst van moderne moleculair-biologische technieken hebben de weg naar de experimentele evolutieleer vrij gemaakt. Vele voorbeelden van evolutionaire adaptatie worden tegenwoordig in detail bestudeerd in modelorganismen, zoals het resistent worden van pathogene bacteriën tegen antibiotica, of de adaptatie van het HIV-virus aan antivirale middelen. Dergelijke studies hebben duidelijk gemaakt dat microben met hun generatieperioden van enkele tientallen minuten zich zeer snel kunnen aanpassen door mutatie en recombinatie van hun genenrepertoire. De toepassing van structuurbepalingstechnieken zoals eiwitkristallografie (Figuur 1) en nucleaire magnetische resonantie (NMR) hebben de moleculaire basis van evolutionaire adaptatie blootgelegd, waarin mutaties in een gen directe structurele veranderingen kunnen

126 veroorzaken in het door het gen gecodeerde genproduct. Selectie en proliferatie van gemuteerde eiwitcomponenten kan zo leiden tot veranderingen in hun activiteit en functie in een cel, wat daarmee de basis vormt voor evolutie op moleculaire schaal.

Figuur 1. Eiwitstructuurbepaling met behulp van röntgendiffractie. Met de klok mee: een eiwitkristal (0,2 x 0,2 x 0,3 mm) gefotografeerd door een lichtmicroscoop, een diffractiepatroon van een eiwitkristal, waarin de intensiteit van de stippen informatie bevat over de structuur van het gekristalliseerde eiwit, en een hogeresolutie elektronendichtheidsmap berekend uit de diffractiedata, waarin de contouren de ruimtelijke verdeling van elektronen in het kristal aangeven.

Dehalogenering van antropogene gehalogeneerde koolwaterstoffen door bacteriën

Het feit dat sommige geselecteerde bodembacteriën kunnen groeien op xenobiotische substraten; chemische stoffen waarvan aangenomen wordt dat zij niet op natuurlijke wijze in de natuur voorkomen, is mogelijk een intrigerend voorbeeld van een heel recente adaptatie. Over de hele wereld zijn verschillende bacteriën geïsoleerd die verschillende gehalogeneerde koolwaterstoffen (Figuur 2), zoals de industriële oplosmiddelen 1,2-dibromoethaan en 1,3-dichloropropanol, als primaire koolstofbron gebruiken. Hoewel veel gehalogeneerde koolwaterstoffen een natuurlijke oorsprong

127 hebben, wordt er van een aantal aangenomen dat zij niet in hoge concentraties voorkwamen voordat ze massaal in de industrie werden gebruikt. Omdat verschillende antropogene gehalogeneerde koolwaterstoffen (geno)toxische eigenschappen bezitten, mag worden verondersteld dat zij een bepaalde evolutionaire druk op bodembacteriën uitoefenen. Het mag dan ook geen grote verrassing heten dat verscheidende bacteriestammen zijn geïsoleerd uit vervuilde grond- en watermonsters die enzymen bevatten die een binding tussen koolstof- en een halogeenatoom kunnen verbreken en het halogeenatoom omzetten in een veel minder toxisch chloride ion. De biochemische en structurele karakterisatie van verschillende dehalogenases in de afgelopen tien jaar heeft gedetailleerd inzicht verschaft in hun katalytisch mechanisme en in een enkel geval hun mogelijke evolutionaire oorsprong.

Figuur 2. Verschillende gehalogeneerde koolwaterstoffen en hun antropogene oorsprong.

Verschillende manieren om een koolstof-halogeen-binding te verbreken

De biochemische karakterisering van verschillende dehalogenases uit uiteenlopende bacteriesoorten heeft duidelijk gemaakt dat verschillende chemische strategieën in de natuur worden gebruikt om de dehalogenering van een substraat mogelijk te maken. Sommige anaërobe bacteriën bevatten cofactor-afhankelijke reducerende dehalogenases die het mogelijk maken om tetrachloroetheen in plaats van zuurstof als elektronenacceptor in de ademhalingsketen te gebruiken. Aërobe bacteriën kunnen cofactor-afhankelijke oxiderende enzymen bevatten die kleine haloalkenen, haloalkanen en sommige aromatische verbindingen kunnen dehalogeneren. Gedetailleerde structurele

128 informatie over deze enzymen is echter nog niet voorhanden, zodat hun werking nog niet volledig duidelijk is. De best gekarakteriseerde dehalogenases waarvan de moleculaire structuren bekend zijn (Figuur 3), maken gebruik van een tweestaps SN2 substitutie mechanisme waarin een tijdelijk covalent intermediair wordt gevormd gebonden aan een aspartaat van het enzym, gevolgd door de hydrolyse van dit intermediair door een water molecuul (Figuur 4A & B). Haloalkaan dehalogenase, haloazijnzuur dehalogenase en de 4- chlorobenzoyl-coenzyme A dehalogenase maken alledrie gebruik van dit mechanisme, maar hebben verschillende aminozuurvolgordes en drie-dimensionale structuren. Dit wijst erop dat zij ontstaan zijn uit evolutionair ongerelateerde superfamilies van enzymen met fundamenteel verschillende activiteiten. Subtiele veranderingen in de actieve holte van de dehalogenases maken de dehalogenase activiteit mogelijk, maar het gebrek aan identiteit in de aminozuurvolgordes met de andere leden van de superfamilies maken onmogelijk om de waarschijnlijke precursor enzymen aan te geven.

Figuur 3. De structuren van verschillende bacteriële dehalogenases. Van links naar rechts: haloalkaan dehalogenase, haloazijnzuur dehalogenase, 4-chlorobenzoyl-coenzym A dehalogenase, haloalcohol dehalogenase, en trans-3-chloroacrylaat dehalogenase.

Moleculaire grondslagen voor twee nieuwe dehalogeneringsactiviteiten

Dit proefschrift beschrijft de opheldering van de moleculaire structuren en katalytische mechanismen van twee evolutionair niet verwante dehalogenases, de

129 haloalcohol dehalogenase HheC uit Agrobacterium radiobacter AD1 en de trans-3- chloroacrylaat dehalogenase CaaD uit Pseudomonas pavonaceae 170. Structuren van de natieve enzymen, en van de enzymen met gebonden producten, substraatanalogen of inhibitors bieden structurele grondslagen voor twee nieuwe dehalogeneringsmechanismen die fundamenteel verschillen van de boven beschreven mechanismen van haloalkaan-, haloazijnzuur-, en 4-chlorobenzoyl-coenzym A dehalogenases. Hoofdstuk 2 geeft een beknopt overzicht van de structuren en reactiemechanismen van de verschillende bacteriële dehalogenases (Figuur 3), waarbij de nadruk ligt op de overeenkomsten en verschillen met structureel verwante enzym families, en hun mogelijke evolutionaire oorsprong. In hoofdstukken 3, 4 en 5 worden de kristallisatie, de structuuropheldering en de werking van haloalcohol dehalogenase HheC beschreven (Figuur 3). HheC speelt een rol in de degradatie van haloalcoholen in A. radiobacter AD1. Het enzym zet haloalcoholen zoals 1,3-dichloro-2-propanol via een eenstaps intramoleculair substitutiemechanisme om in het overeenkomstige epoxide en HCl (Figuur 4C). De resultaten laten zien dat HheC verwant is aan de zogenaamde korte dehydrogenases/reductases (SDRs), een familie van NAD(P)H-afhankelijke redox enzymen die in bijna alle organismen voorkomen. Structuren van HheC met gebonden dehalogeneringsprodukten en een haloalcohol analoog laten zien dat de dehalogeneringsreactie mogelijk wordt gemaakt door een geconserveerde Ser-Tyr-Arg katalytische triade. De bindingsplaats van NAD(P)H ontbreekt, maar deze is vervangen door een halidebindingsplaats die het cofactor- onafhankelijke dehalogeneringsmechanisme mogelijk maakt. Structurele vergelijking met verschillende bekende structuren van SDR enzymen laat overtuigend zien dat de zes haloalcohol dehalogenases die op dit moment bekend zijn waarschijnlijk zijn voortgekomen uit twee verschillende NADH-bindende, in plaats van NADPH-bindende, SDR enzymen. Hoofdstuk 6 beschrijft de moleculaire structuren van natief trans-3-chloroacrylaat dehalogenase CaaD (Figuur 3), en van het enzym dat geïnactiveerd is door inhibitor die het eiwit covalent modificeert. Het enzym is instaat een afbraakproduct van het veelgebruikte bestrijdingsmiddel 1,3-dichloropropeen, trans-3-chloroacrylaat, te dehalogeneren. De resultaten bieden gedetailleerd inzicht in een nieuw dehalogeneringsmechanisme, waarin een koolstof-chloorbinding in trans-3-chloroacrylaat via een Michael-additie van water wordt verbroken. Het product van deze additie reactie is zeer instabiel, en valt vervolgens spontaan uit elkaar in malonaatsemialdehyde en HCl. CaaD is gerelateerd aan de familie van 4-oxalocrotonaat tautomerases (4-OTs). 4OTs

130 gebruiken een N-terminale proline (Pro1) als een katalytische base in tautomerisatie reacties, terwijl CaaD de proline als een zuur gebruikt. Een naburige glutamaat die niet aanwezig is in de tautomerases fungeert als een water-activerende katalytische base, en maakt samen met de proline de hydratase activiteit van CaaD mogelijk.

Figuur 4. Reactiemechanismen van de verschillende dehalogenases. A & B) Schematische weergave van de substitutie stap gekatalyseerd door een nucleofiele aspartaat in haloalkaan-, haloazijnzuur-, en 4-chlorobenzoyl-coenzym A dehalogenase. De substitutie van het halogeen atoom produceert in alle gevallen in een covalent intermediair, dat vervolgens wordt gehydrolyseerd door een water molecuul. C) Schematische weergave van de intramoleculaire substitutiereactie zoals die gekatalyseerd wordt door haloalcohol dehalogenase HheC. D) Schematische weergave van de Michael additie van water aan de dubbele binding in trans-3-chloroacrylaat door trans-3- chloroacrylaat dehalogenase CaaD, wat een instabiel haloalcohol product oplevert.

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De oorsprong van dehalogeneringsactiviteiten in bodembacteriën

Hoewel de evolutionaire oorsprong van de meeste gekarakteriseerde dehalogenases onduidelijk blijft, biedt de structurele informatie over haloalcohol dehalogenase HheC en trans-3-chloroacrylaat dehalogenase inzicht in hun mogelijke evolutionaire oorsprong. In tegenstelling tot de haloalkaan- en haloazijnzuur dehalogenases, zijn er maar een paar haloalcohol dehalogenase en chloroacrylaat dehalogenase sequenties bekend. Hun zeldzame voorkomen in speciaal geselecteerde bodembacteriën, en hun ontbreken in andere organismen, suggereert dat zij mogelijk recent geëvolueerd kunnen zijn uit gerelateerde enzymen onder druk van haloalcoholen in hun leefomgeving. Katalytische promiscuïteit van een precursor enzym kan hierbij een grote rol hebben gespeeld. Een SDR precursor van de haloalcohol dehalogenases kan toevallig een haloalcoholsubstraat hebben kunnen binden, waarbij het halogeenatoom mogelijk gebonden werd in de NADH bindingsplaats. De katalytische tyrosine zou vervolgens de hydroxylgroep van de alcohol hebben kunnen deprotoneren, en daarmee de intramoleculaire substitutiereactie hebben kunnen katalyseren. Zo’n promiscue dehalogenase activiteit kan vervolgens evolutionair zijn geoptimaliseerd. Een dergelijk scenario is ook aannemelijk voor het ontstaan van chloroacrylaat dehalogeneringsactiviteit. In tegenstelling tot de SDR enzymen, is voor enkele tautomerases een dergelijke katalytische promiscuïteit al experimenteel aangetoond. Eén van de grote wetenschappelijke uitdagingen voor de toekomst zal zijn om deze hypothesen te verifiëren met behulp van experimentele evolutie.

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Het groene licht is gegeven door de leescommissie bestaande uit de professoren Jan Engberts, Dick Janssen en Alan Mark, waarvoor ik hen hartelijk bedank. Na iets meer dan vier jaar onderzoek in de eiwitkristallografie groep van Prof. Bauke Dijkstra is het nu tijd dit proefschrift af te maken door het schrijven van dit nawoord. Het was een leuke, intensieve en vooral leerzame tijd die al begon als student, toen ik wegwijs werd gemaakt in de wereld van de bioinformatica en eiwitstructuurmodeling door Milou Kouwijzer en Rob van Montfort. Het door Bauke aangeboden promotieproject, de structuuropheldering van een enantioselectieve haloalcohol dehalogenase, sprak me erg aan, mede door de beoogde intensieve samenwerking met biochemici uit de groep van Prof. Dick Janssen. Het was een luxe om met Bauke, Dick, Johan, Marco, Lixia, Jeffrey, Erik, en later ook Hanya, mijn enthousiasme voor dit project te kunnen delen. Jan Tiesinga kwam als een geschenk uit Leiden om een jaar enthousiast met me mee te werken, en werd later een collega AIO in de groep. Met de klonering en zuivering van een 3-chlooracrylzuurdehalogenase door Gerrit Poelarends, eerst AIO in de groep van Dick Janssen, later post-doc in de groep van Prof. Chris Whitman aan de Universiteit van Texas, kreeg ik de kans om nog een ander dehalogenerend enzym te kristalliseren. Gerrit en Chris leverden het eiwit en de covalent-bindende remmers die het enzym onomkeerbaar inactiveerden. Wim Brugman ging als student hiermee aan de slag, en ondanks de moeilijke kristallisatie lukte het ons vlak voor zijn afstuderen de struktuur op te lossen. De succesvolle afloop van deze onderzoeksprojecten heb ik dan ook voor een deel aan de bovenstaande mensen te danken. Bij de start van een onderzoeksproject in een biofysisch lab kan je niet zonder de ervaren collegas om je heen, want zij hebben (meestal) een antwoord op je vele vragen. Allesanda, Andy, Arjan, Arthur, Charles, Els, Erik, Fabrizia, Francesca, Frans, Gertie, Gerrit Groenhof, Gregory, Henriëtte, Jan Drenth, Joost, Kor, Marc Lensink, Marco, Rob, Roberto, mijn paranimf Thomas, Tjaard, en Wilco, bedankt voor alle tips ’n tricks die ik in de loop van de jaren van jullie heb gekregen. Ook alle mensen die na mij in de groep arriveerden, Andre, Andrea, Avinash, Cyril, Hiromi, Jan, Karin, Wangsa, wil ik hierbij bedanken voor de gezellige tijd. Ik hoop dat jullie op het lab ook wat aan mij hebben gehad. Ook buiten het lab heb ik mijn omgeving in Groningen als uiterst rijk en inspirerend ervaren. Met de mensen van stichting Holodeck, A3, Dennis, Dinanda, Jachim, Jeff, Jullius, Kim, Linda, Marije, Martijn, Nelleke, Renske, Ruben, Simon en Wouter, was het een fijne afwisseling om samen te werken aan zoiets radikaal anders als de ClubPlus feesten. Ook vele vrienden en kennissen uit het dag- en nachtleven van Groningen, in het bijzonder Jaap Bouma, Gideon van der Staaij, Maarten Pragt, mijn tweede paranimf Peter Krako, Peter Kremer, Rienk Eelkema en Rop Broere, wil ik hier bedanken voor een fijne tijd. Tot slot bedank ik mijn ouders en zus voor hun onvoorwaardelijke steun door de jaren heen, en mijn lieve vriendin Debby, die de vele euforische, maar ook de moeilijkere momenten tijdens het ontstaan van dit proefschrift van zeer nabij heeft meegeleefd.

René

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Stellingen Stellingen behorend bij het proefschrift behorend bij het proefschrift “Molecular basis of two novel dehalogenating activities. “Molecular basis of two novel dehalogenating activities. Structure/function relationships in an evolutionary context” Structure/function relationships in an evolutionary context” René M de Jong, Groningen, 30 januari 2004 René M de Jong, Groningen, 30 januari 2004

1. Het meenemen van structurele informatie in het uitlijnen van aminozuurvolgordes kan 1. Het meenemen van structurele informatie in het uitlijnen van aminozuurvolgordes kan verrassende nieuwe inzichten opleveren. verrassende nieuwe inzichten opleveren. Hoofdstuk 4 van dit proefschrift. Hoofdstuk 4 van dit proefschrift.

2. Het clichébeeld van de wetenschapper als specialist met enkel notie voor zijn eigen 2. Het clichébeeld van de wetenschapper als specialist met enkel notie voor zijn eigen discipline wordt bevestigd door het ongebreideld voortbestaan van al lang discipline wordt bevestigd door het ongebreideld voortbestaan van al lang achterhaalde onjuistheden in de literatuur rond het natuurlijk voorkomen van het achterhaalde onjuistheden in de literatuur rond het natuurlijk voorkomen van het chlooratoom. chlooratoom. Oberg, G (2002): The natural chlorine cycle - fitting the scattered pieces. Appl. Microbiol. Biotechnol. 58, 565- Oberg, G (2002): The natural chlorine cycle - fitting the scattered pieces. Appl. Microbiol. Biotechnol. 58, 565- 581. 581.

3. Het ontlast het geweten te beseffen dat zeer milieuonvriendelijke en toxische 3. Het ontlast het geweten te beseffen dat zeer milieuonvriendelijke en toxische gehalogeneerde verbindingen als dioxines naast een industriële oorsprong, ook een gehalogeneerde verbindingen als dioxines naast een industriële oorsprong, ook een preïndustriële antropogene oorsprong en een natuurlijke oorsprong hebben. preïndustriële antropogene oorsprong en een natuurlijke oorsprong hebben. Meharg, AA & Killham, K (2003): A pre-industrial source of dioxins and furans. Nature 421, 909-910. Hoekstra Meharg, AA & Killham, K (2003): A pre-industrial source of dioxins and furans. Nature 421, 909-910. Hoekstra et al. (1999): Natural formation of chlorinated phenols, dibenzo-p-dioxins, and dibenzofurans in soil of a et al. (1999): Natural formation of chlorinated phenols, dibenzo-p-dioxins, and dibenzofurans in soil of a Douglas fir forest. Env. Sci. Technol. 33, 2543-2549. Alcock, RE & Jones, KC (1996): Dioxins in the Douglas fir forest. Env. Sci. Technol. 33, 2543-2549. Alcock, RE & Jones, KC (1996): Dioxins in the environment: a review of trend data. Env. Sci. Technol. 30, 3133-3143. environment: a review of trend data. Env. Sci. Technol. 30, 3133-3143.

4. Het ontbreken van een halidebindingsplaats in een dehalogenerend enzym is geen 4. Het ontbreken van een halidebindingsplaats in een dehalogenerend enzym is geen goed argument voor de speculatie dat de dehalogeneringsreactie een “toevallige” goed argument voor de speculatie dat de dehalogeneringsreactie een “toevallige” zijreactie van een hydraterend enzym was, die vervolgens gerekruteerd zou zijn om zijreactie van een hydraterend enzym was, die vervolgens gerekruteerd zou zijn om als dehalogenase te functioneren. als dehalogenase te functioneren. Wang, S et al. (2003): Reactions of trans-3-chloroacrylic acid dehalogenase with acetylene substrates: Wang, S et al. (2003): Reactions of trans-3-chloroacrylic acid dehalogenase with acetylene substrates: consequences of and evidence for a hydration reaction. Biochemistry 42, 8762-8773. consequences of and evidence for a hydration reaction. Biochemistry 42, 8762-8773.

5. Het wordt tijd voor een opstand van intellectuelen tegen de debilisering van de 5. Het wordt tijd voor een opstand van intellectuelen tegen de debilisering van de maatschappij. maatschappij. Bart Nooteboom, NRC Handelsblad, 1 mei 2001 Bart Nooteboom, NRC Handelsblad, 1 mei 2001

6. Dat Tiësto zich al twee jaar de beste DJ ter wereld mag noemen, is tekenend voor het 6. Dat Tiësto zich al twee jaar de beste DJ ter wereld mag noemen, is tekenend voor het commerciële hoogtepunt, maar ook het artistieke dieptepunt waarop de hedendaagse commerciële hoogtepunt, maar ook het artistieke dieptepunt waarop de hedendaagse dance-cultuur zich bevindt. dance-cultuur zich bevindt.

7. De ontwikkeling van underground techno en house tot mondiaal gewaardeerde 7. De ontwikkeling van underground techno en house tot mondiaal gewaardeerde muziekstromingen die niet worden gehinderd door taalbarrières en cultuurverschillen, muziekstromingen die niet worden gehinderd door taalbarrières en cultuurverschillen, is een vorm van culturele globalisering. is een vorm van culturele globalisering.

8. Het massale gebruik van het Windows besturingssysteem met haar vele 8. Het massale gebruik van het Windows besturingssysteem met haar vele beveiligingsproblemen maakt de strikte geheimhouding van de broncode een poging beveiligingsproblemen maakt de strikte geheimhouding van de broncode een poging tot ondermijning van de internationale veiligheid. tot ondermijning van de internationale veiligheid.

9. Een verbod aan koffieshops om aan buitenlanders te verkopen zal door een explosief 9. Een verbod aan koffieshops om aan buitenlanders te verkopen zal door een explosief toenemende illegale handel van Nederland pas echt een narcostaat maken. toenemende illegale handel van Nederland pas echt een narcostaat maken.

10. Voor alles is een norm. Jammer genoeg hecht niet iedereen daar waarde aan. 10. Voor alles is een norm. Jammer genoeg hecht niet iedereen daar waarde aan.