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Divergent or just different Rozeboom, Henriette

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Divergent or just different Structural studies on six different enzymes

Henriëtte Rozeboom

Printed by Ipskamp Drukkers, Enschede

The research presented in this thesis was carried out in the Crystallography group at the Groningen Biomolecular Sciences and Biotechnology Institute. Faculty of Mathematics and Natural Sciences, University of Groningen, the Netherlands.

Cover: The image on the front shows the Glucan complexed with the inhibitor 5-fluoro-idopyranose. The images on the back show on the left bottom Naproxen esterase and right bottom CesB. The top left image shows Mannuronan C5 epimerase complexed with the substrate mannuronan trisaccharide and the right top image shows the glycosylated endo-xylogalacturonan .

ISBN (print version) 978-90-367-7337-9 ISBN (electronic version) 978-90-367-7336-2

Divergent or just different

Structural studies on six different enzymes

Proefschrift

ter verkrijging van de graad van doctor aan de Rijksuniversiteit Groningen op gezag van de rector magnificus prof. dr. E. Sterken en volgens besluit van het College voor Promoties.

De openbare verdediging zal plaatsvinden op

vrijdag 14 november 2014 om 14.30 uur

door

Henriëtte Jantiena Rozeboom geboren op 12 juli 1964 te Appingedam

Promotor

Prof. dr. B.W. Dijkstra

Beoordelingscommissie

Prof. dr. D.B. Janssen Prof. dr. G.J. Poelarends Prof. dr. T.M. Sixma

Voor pap en mam♥, Cees, Lisa en Bo Wan

Contents ......

Page

1 General introduction 9 ......

Structural and mutational characterization of the catalytic A- 37 2 module of the mannuronan C-5-epimerase AlgE4 from Azotobacter vinelandii ...... Crystal structure of α-1,4-glucan lyase, a unique glycoside 59 3 hydrolase family member with a novel catalytic mechanism ......

Crystal structure of quinone-dependent alcohol 85 4 dehydrogenase from Pseudogluconobacter saccharoketogenes - A versatile dehydrogenase oxidizing alcohols and carbohydrates ...... Crystal structure of endo-xylogalacturonan hydrolase from 105 5 Aspergillus tubingensis ......

Crystal structures of two Bacillus carboxylesterases with 119 6 different enantioselectivities ......

7 Summary, General discussion & Outlook 141 ......

8 Nederlandse samenvatting 149 ......

References 155 CV & Publications 173 Dankwoord 177

7

8

Chapter 1

General introduction

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ABSTRACT

In this introductory chapter we have chosen to focus on an evolutionary view of protein structure and function. Such a view is of interest as it gives us insight into the inner workings of the evolutionary processes at the molecular level. Understanding how enzymes evolved as biological catalysts to perform the wide variety of reactions found across all kingdoms of life is fundamental to a broad range of biological studies. The evolutionary process is illustrated with seven examples of enzymes/functional . The three-dimensional structures of these proteins and their evolutionary relationships to other proteins and enzymes are discussed. Furthermore, the technique, X-ray crystallography, by means of which these structures have been determined, is shortly reviewed.

1. BIOLOGICAL EVOLUTION

About 4.6 billion years ago Earth was formed and soon after, 3.6 billion years ago, primordial life began to evolve and the first simple cells appeared (Schopf et al., 2007). How life arose on Earth is still a great mystery. The hypothesis is that life started in a primitive “RNA World” and that later DNA and proteins came into existence (Joyce, 2002). Today, it is generally assumed that the last universal common ancestor of all current living organisms was most likely a single-celled organism that lived ~3.5 billion years ago (Schopf et al., 2007, Woese, 1998). Since then cellular life has split and evolved into the modern life forms belonging to the three ‘domains’ of life we now know as Archaea, Bacteria and Eukaryota. An important event in the early evolution of life was the emergence of oxygenic photosynthesis, which evolved only once, about 2.4 billion years ago, in a common ancestor of current cyanobacteria that emitted O2 as a waste product (Schirrmeister et al., 2013). Chloroplasts found in eukaryotes (algae and plants) likely evolved from an endosymbiotic relation with such a cyanobacterial ancestor about 1 billion years ago.

Although initially the emitted molecular oxygen (O2) was captured by dissolved iron and organic matter, later O2 started to accumulate in the atmosphere. The increasing oxygen concentration forced some obligate anaerobic organisms to adapt, and oxygen- dependent life forms, including multicellular organisms, evolved (Schirrmeister et al., 2013). Nowadays, almost all life on Earth depends ultimately on photosynthetic primary producers, like cyanobacteria and plants (Sleep et al., 2008). Evolution spans billions of years. Tracing the early evolution of life, by fossil dating can generate clues on the origin of a species. However, identifiable fossil bacteria are not particularly widespread. In addition, it is often very difficult to obtain information about

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General introdution ......

(extinct) common ancestors. For instance, it is impossible to go far back in time to sequence proteins or DNA from far ancestors that, by definition, no longer exist. In practice, we may bridge a gap of about 100,000-150,000 years as shown by the successful sequencing of the entire genome extracted from the toe bone of a 130,000- year-old Neanderthal woman found in a Siberian cave. According to the genome sequence the Homo neanderthalensis is only 0.3% different in its coding sequences from Homo sapiens (Prufer et al., 2014). This research and more to come might shed light on the mystery as to how we evolved and why modern humans are the only surviving human lineage.

2. PROTEIN EVOLUTION

Nowadays, it is generally assumed that all currently existing proteins diverged from a rather small set of proteins (a few hundreds, possibly ~1000), which already existed in the last universal common ancestor (Elias et al., 2012). The earliest enzymes were probably generalists, capable of catalyzing particular chemical transformations with broad substrate specificity. During evolution new enzymes emerged by incremental , gene duplications, gene fusion, gene recruitment, gene transfer and post- translational modification (Chothia et al., 2009). Gene duplication provides new genetic material for increasing protein repertoires. The duplicated gene sequence (a paralogue) diverges by mutations, deletions, and insertions to produce modified proteins that may have useful new properties (Chothia et al., 2003). One copy of the gene continues to produce proteins that keep their original, critical function, while the other copy can evolve independently, producing proteins with new and different functions. In general, evolution leads to sequence and structural divergence, which benefits survival at the level of cells and organisms. An important question concerns the rate of evolution. Is it possible to determine how many years ago two organisms (or proteins) diverged? The molecular clock hypothesis proposes that the rate of evolution of a given protein (or, later, DNA sequence) is approximately constant over time and evolutionary lineages (Zuckerkandl et al., 1965, Morgan, 1998). More specifically, the hypothesis proposes that a statistical correlation exists between the time elapsed since the last common ancestor of two contemporary homologous proteins and the number of amino acid differences between their sequences. In practice, this would allow biologists to give a temporal dimension to phylogenetic trees (see below) constructed from molecular data (Morgan, 1998). However, determining the absolute timing of events using molecular clocks has attracted severe skepticism for several reasons. Different proteins appear to evolve at different rates, differences in generation times of organisms may affect the rate, and the size of the population is important as well (in small populations more mutations are neutral)

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(Ayala, 1999). All these factors make strict molecular clocks less useful for predictions. Instead, a “relaxed molecular clock” has been proposed that is better able to co-estimate phylogeny and divergence times (Drummond et al., 2006). Another model of evolutionary rate is the “Universal Pacemaker” model (Snir et al., 2012), which is based on the observation that when the rate of evolution changes, the change occurs synchronously in many if not all genes in an evolving genome. These more general models may overcome (partly) the limitations of the molecular clock hypothesis. Although the rate of mutational change during evolution is, in general, not very precisely defined, the similarities/relationships between organisms, proteins, or DNA sequences can be quantified much more easily. Such evolutionary relationships are often shown in a phylogenetic tree or evolutionary tree, first sketched by Darwin in his notebook in 1837 (Ragan, 2009). The phylogenetic tree is based upon similarities and differences in the external features and/or genetic characteristics of a certain group of related biological species or gene/protein sequences (taxa). The branches joined together in a node of the tree are implied to have descended from a common ancestor (Drummond et al., 2006).

Figure 1. Example of a phylogenetic tree of PQQ-dependent alcohol dehydrogenases obtained with the maximum likelihood method as implemented in MEGA6 (Tamura et al., 2013).

2.1 Evolution and protein folds

The first crystal structures of (Kendrew et al., 1958) and, a few years later, of (Perutz et al., 1960) showed that the conformations of the myoglobin and hemoglobin peptide chains are very similar (Cullis et al., 1962, Huber et al., 1971). Although the sequence identity of these two is only 24%, they have a striking structural similarity as indicated by their root mean square difference (RMSD) value (see below) of only 1.6 Å. Thus, their 3D structures are much better conserved than their amino acid sequences. The better conservation of 3D structures compared to amino acid

12

General introdution ...... sequences has later been confirmed for many other protein families (Chothia et al., 1986). Indeed, detecting the evolutionary relationships of proteins from sequence comparisons can be very difficult, especially if their sequences have diverged beyond recognition. In contrast, 3D structures diverge far less rapidly than sequences. Restraints imposed on the tertiary structure of the protein preserve a close-packed hydrophobic interior and conserve the functional properties of the (Chothia et al., 1987). Therefore, evolutionary relationship of proteins can often be recognized from their 3D structures, even if the primary structures do not give clear clues (Murzin, 1998). Similarities in sequence and 3D structure have allowed the classification of proteins. Proteins with significant sequence homology are clustered in a family; different families with common structure and function are grouped into protein superfamilies. Classification of proteins into superfamilies is detailed in databases such as Structural Classification Of Proteins (SCOP or SCOP2; (Murzin et al., 1995, Andreeva et al., 2014)) and Class Architecture Topology and Homology (CATH; (Sillitoe et al., 2013)). The large amount of structural data coming from structural genomics projects greatly increases the ability to classify protein domains in terms of their evolutionary relationships (Teichmann et al., 2001). The extent of the structural divergence can be quantified by rigid body superposition of the common cores of homologous proteins and calculating the root mean square differences in the positions of their main-chain or Cα atoms (Chothia et al., 1986).

2.2 Divergent evolution

Divergent evolution is the evolutionary development of proteins that differ from each other in form and function but that have evolved from the same common ancestor and that usually belong to the same superfamily (Galperin et al., 2012). This type of evolution is most extensively described in the literature and we have chosen to discuss three classic examples of divergent evolution, i.e. the family (myoglobins and ), lysozymes, and crystallins.

2.2.1 The globin family

As shown above, the structure of hemoglobin revealed a close structural similarity to the structure of myoglobin. Both proteins belong to the very large globin . Globin family members are believed to have evolved from a common ancestor. A detailed analysis of the myoglobin and hemoglobin crystal structures by Lesk and Chothia revealed that mutations of buried residues cause rigid-body movements of the α-helices relative to each other and associated changes in the turn regions, but that the geometry

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Chapter 1 ......

of the active site is not affected (Lesk et al., 1980). Later, it emerged that nearly all residues in the globin peptide chain can be mutated without compromising the active site (Bashford et al., 1987). This shows the power of evolution: only functional variants are retained. On the other hand, the requirement that function is retained puts severe limitations to the development of new functionalities. It is believed that two whole- genome duplication events (during meiosis) in the globin gene superfamily in the stem lineage of vertebrates were necessary for promoting evolutionary innovation (Storz et al., 2013). Hemoglobin and myoglobin are good examples of divergent evolution, both probably having arisen from a proto-myoglobin (Aguileta et al., 2006). The single-chain storage protein myoglobin has only one group and serves as a reserve for oxygen in muscle and other tissue. The major hemoglobin in adult humans, (HbA), is a heterotetramer composed of two α-globin and two β-globin polypeptides, each with an associated heme group, which transports oxygen to the cells. The tetrameric organization of hemoglobin tetramer is crucial for its allosteric and oxygen-delivery properties, and the presence of two different globin chains in the tetramer is essential for this function. The α-globin and β-globin chains (42 % sequence identity) are believed to have arisen from the same primordial hemoglobin gene by a gene duplication event, with afterwards each gene following an independent evolutionary history. Other variants of hemoglobin are hemoglobin A2 (HbA2), and F (HbF). The latter hemoglobin consists of two α- and two γ-globin polypeptide chains, which differ at 39 positions from β-globin; HbF has a higher affinity for oxygen because of weaker subunit interface interactions. Moreover, HbF binds the hemoglobin regulator 2,3- biphosphoglycerate less tightly (Frier et al., 1977). These properties ensure delivery of oxygen to the growing fetus. HbA2 is found in trace amounts in the blood and has two α- globins and two δ-globins, which differ at 10 positions from β-globin and which are more resistant to thermal denaturation than HbA. δ-Globins inhibit the polymerization of sickle cell deoxyhemoglobin (HbS) (Sen et al., 2004). γ-Globin and δ-globin evolved from an ancestral β-type globin by further gene duplication. The different isoforms of hemoglobin, resulting from divergent evolution, function according to different tissue requirements for oxygen. Additional heme-containing globins, like and , have been discovered by data mining (Hardison, 2012), but their functional roles are not yet fully clear. Furthermore, plants also contain hemoglobins called , which, in root nodules, are involved in oxygen transport to symbiotic nitrogen-fixing bacteria or are used as oxygen scavengers/sensors (Hoy et al., 2007).

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2.2.2 Lysozyme

Lysozymes are defence enzymes mainly found in egg whites, tears and various secretions of eukaryotic cells. In 1965 hen egg-white lysozyme (HEWL, C (chicken) type) was the very first enzyme of which the atomic structure was solved (Blake et al., 1965). Later, structures were determined of goose type (G type) (Grütter et al., 1983), phage-type (T4) (Matthews et al., 1974) and invertebrate type (I type) (Goto et al., 2007) lysozyme. Matthews & Remington, who solved the 3D structure of T4 phage lysozyme, noticed that its structure was quite different from that of hen egg-white lysozyme (Matthews et al., 1974). However, in 1976 Rossmann & Argos recognized the structural similarity between hen egg-white lysozyme and phage-type lysozyme via rotation and translation superposition of the two structures (Rossmann et al., 1976). This study was the first example of the discovery of an evolutionary relationship of two proteins by comparison of their 3D structures, in the absence of noticeable amino acid sequence homology. In 1994 Thunnissen et al. published the crystal structure of the E. coli lytic transglycosylase SLT70 (Thunnissen et al., 1994). SLT70 appeared to have a C-terminal domain with a lysozyme-like fold, which contained the enzyme’s active site, where the lytic transglycosylase activity resides. Although the amino acid sequence homology between the four representative lysozyme types (C, G, T4, I) and C-SLT70 is very limited, their three-dimensional structures show striking similarities and indicate their evolutionary relationships (Thunnissen et al., 1995a). The lysozyme fold is composed of two domains, separated by a deep cleft containing the active site. One domain mainly consists of a β-sheet structure, while the other domain is more helical in nature (Callewaert et al., 2010). The conserved structural core of the lysozyme fold comprises five secondary structure elements that are arranged in a topologically identical order around the substrate- binding cleft. The secondary structure elements comprise a three-stranded β-sheet and two α-helices in the N-terminal lobe and two small α-helical regions in the C-terminal lobe, but with somewhat different orientations in the various proteins (Thunnissen et al., 1995a), similar to what was observed for the globin family (Lesk et al., 1980). Yet, the substrate-binding cleft (where peptidoglycan binds) and part of the active site have been conserved during evolution. In contrast, the catalytic residues are not fully conserved. Whereas C and I type lysozymes contain two catalytic acidic amino acids in their active site — the acid/base Glu and an Asp that serves as a nucleophile —, in G-type lysozyme and C-SLT70 only the Glu is present. In T4 lysozyme the acid/base catalyst Glu is conserved as well, but this enzyme has also an Asp, which is however not essential for catalysis, and which is at a different position compared to the nucleophilic Asp in C-type lysozyme (Kuroki et al., 1999). These differences clearly suggest that not only the folds of the enzymes have diverged, but also the catalytic mechanisms. Rather than having an enzyme-based

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Chapter 1 ......

nucleophile, T4 and G-type lysozyme, as well as C-SLT70, make use of the substrate’s N- acetyl group to stabilize the reaction intermediate (van Heijenoort, 2011). In SLT70 even the reaction specificity has changed: rather than catalyzing a hydrolysis reaction as the lysozymes do, it catalyzes a 1,6)-transglycosylation reaction via an intramolecular nucleophilic attack. These examples show that during evolution the substrate-binding sites and the catalytic Glu remained conserved, but that other solutions emerged for the stabilization of the oxocarbenium ion reaction intermediate. Interestingly, α-lactalbumin, a protein expressed in the lactating mammary gland, has also a lysozyme-like fold, resembling in particular calcium-binding C-type lysozymes found only in a few species of birds and mammals, with which it shares about 35-40% amino acid sequence identity (baboon and human) and four conserved disulfide bridges (Acharya et al., 1989). In α-lactalbumin a Tyr residue blocks the substrate-binding cleft and the catalytic acid/base Glu is not present (Acharya et al., 1989). Gene duplication of a common ancestor and subsequent divergent evolution are thought to be the evolutionary events leading to the coexistence of lysozymes and α-lactalbumins in mammals. Although both proteins have anti-infective activity in common, their functions are quite distinct (Callewaert et al., 2010, Qasba et al., 1997). Whereas lysozymes degrade the bacterial cell wall polymer peptidoglycan, α-lactalbumin is the regulatory subunit of the lactose synthase complex, which is required for the synthesis of lactose in milk. The C-type lysozyme and α-lactalbumin structures represent a clear example of functional divergence preceding gross structural divergence.

Figure 2. Cartoon representation of the crystal structures of (A) the common fold of lysozymes with the two N- terminal lobe helices in magenta, the C-terminal lobe helix in brown, the 3-stranded β-sheet in cyan; the catalytic glutamate and aspartate are depicted as sticks, the green spheres are the calcium ions, (B) Gallus gallus (hen egg- white, C type) lysozyme (PDB ID 2LYZ (Blake et al., 1965, Diamond, 1974), (C) Tachyglossus aculeatus (Echidna milk) calcium containing lysozyme (PDB ID 1JUG (Guss et al., 1997)), (D) Papio cynocephalus (baboon milk) α- lactalbumin (PDB ID 1ALC (Acharya et al., 1989)), (E) Bacteriophage T4 lysozyme (PDB ID 2LZM (Weaver et al., 1987), (F) Anser anser (G type) lysozyme (PDB ID 153L (Weaver et al., 1995), (G) Escherichia coli soluble lytic transglycosylase C-terminal domain (PDB ID 1SLY (Thunnissen et al., 1995b)and (H) Tapes japonica (Japanese cockle, I type) lysozyme (PDB ID 2DQA, (Goto et al., 2007) (for easy viewing the B subunit is not shown).

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2.2.3 Crystallins

Crystallins are also very interesting from an evolutionary perspective. They are major lens proteins of the vertebrate eye, and in the eye they are closely packed, and need to be extremely long-lived. They also occur in other tissues, where they have different roles, usually being involved in metabolic processes (Slingsby et al., 2013). This phenomenon, that the same protein fulfills different functions in different cells or organs (also called protein moonlighting), involves the acquisition of a novel secondary function. Usually, this involves a change in gene expression without loss of the primary function, or it involves duplication of the gene (Tsai et al., 2005). Structurally, α-crystallins are related to small heat-shock proteins (sHSP), which are expressed in many tissues, notably muscle (Slingsby et al., 2013). Remarkably, the same protein is used as α-crystallin in the lens and has a function as chaperone in other tissues (as well as in the lens). Other examples of enzymes moonlighting as a crystallin include α-/τ-crystallin and argininosuccinate lyase (ASL)/δ2-crystallin (Wistow, 1993, Piatigorsky, 2003). This latter case is of some interest, since in ducks two δ-crystallin isoforms exist, δ1- and δ2-crystallin, which have 94% amino acid sequence identity. While duck δ2-crystallin has maintained ASL activity, evolution has rendered duck δ1- crystallin enzymatically inactive (Tsai et al., 2005). Thus, following the recruitment of ASL as δ-crystallin in an early ancestor of birds and reptiles, adaptive conflict may have provided the selective pressure for gene duplication, allowing one gene to make subtle sequence modifications necessary to enhance lens function, while the other maintained enzymatic function (Wistow, 1993). Similarly, the homologous pair S- crystallin/glutathione S- (GST) has also evolved via gene duplications with subsequent divergence (Piatigorsky, 2003). Evidently, structurally stable enzymes have been recruited as lens proteins several times during evolution (Wistow et al., 1988).

2.3 Convergent evolution

Enzymes that belong to different folds or superfamilies and that that do not show amino acid sequence homology, but that share the same substrate and reaction specificity and that have strikingly similar active site arrangements are commonly described as the outcome of convergent evolution and are referred to as analogous (Elias et al., 2012). A few examples of convergent evolution are described in the next sections; they are of proteins on which substantial structural information was gathered in the Laboratory of Biophysical Chemistry of the University of Groningen.

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2.3.1 Proteases

Convergent evolution of active sites with the same organization of acid-base-nucleophile (Asp-His-Ser) was first identified in the two unrelated serine proteases α-chymotrypsin (Matthews et al., 1967) and subtilisin BPN’. The structure of this latter enzyme was independently determined by Kraut et al. (Wright et al., 1969) and Drenth et al. (Drenth et al., 1971) from crystals obtained from rather different crystallization conditions. Chymotrypsin and subtilisin BPN’ have completely different folds, but both feature an active site with Ser, His and Asp as catalytic residues. Their active sites were compared by Matthews et al. (Matthews et al., 1977), concluding that in both enzymes the Ser acts as the nucleophile in the reaction, and the Asp-His pair as the for a proton in the transition state. Interestingly, in 1968 Jan Drenth and coworkers had solved the 3D structure of papain (Drenth et al., 1968), a cysteine protease, which showed a surprisingly analogous of Cys, His, Asn, but a fold entirely different from that of α-chymotrypsin and subtilisin. Serine proteases have been classified into 13 clans. The three major clans, based on their structures, are chymotrypsin/trypsin-like, subtilisin-like and α/β hydrolase-like (see also below) (Krem et al., 2001). Chymotrypsin-like enzymes have a β/β structure with a sequence order of the catalytic residues of His-Asp-Ser. Subtilisin-like enzymes have a α/β/α structure with the catalytic residues in the order Asp-His-Ser. Finally, α/β (e.g. lipases, esterases and prolyl oligopeptidases) have a mostly parallel - structure flanked by -helices with their catalytic residues in yet another sequence order (Ser-Asp-His). From this short overview it can be concluded that the catalytic triad exists in a variety of forms and evolved independently a number of times (Dodson et al., 1998).

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Figure 3. Catalytic triads of three different serine proteases and a cysteine protease. Nitrogen atoms are shown in blue, oxygen atoms are shown in red, sulphur atoms are shown in yellow, H-bonds are indicated by dotted lines. (A) Bos taurus chymotrypsinogen (PDB ID 1CHG (Freer et al., 1970), (B) Bacillus amyloliquefaciens subtilisin (PDB ID 1SBT (Alden et al., 1971), (C) α/β hydrolase, Pseudomonas aeruginosa lipase (PDB ID 1EX9 (Nardini et al., 2000)and (D) cysteine protease, Carica papaya papain (PDB ID 1PAD (Drenth et al., 1968).

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2.3.2 Hemocyanin - tyrosinase

In contrast to vertebrates, which use hemoglobin as oxygen-carrying protein, some invertebrates deploy hemocyanin to transport oxygen. Hemocyanins are large, blue- colored, multimeric proteins with allosteric oxygen-binding properties similar to hemoglobin (Van Bruggen, 1980). Two different families of hemocyanins exist, arthropodan (e.g. arachnids and crustaceans) and molluscan (such as squid, cuttlefish, octopus, snails and slugs) hemocyanins. The evolutionary relationship between these hemocyanins has long been a topic of speculation: are they diverged gene products or the result of convergent evolution? Both hemocyanin families have very different amino acid sequences, structures and subunit composition, but there is evidence for a common origin of at least part of their active site (Durstewitz et al., 1997). Hemocyanins contain a binuclear type-3 copper site that reversibly binds a single oxygen molecule (O2). This copper site contains two copper ions (Cu-A and Cu-B), each ligated by three histidine residues from different pairs of helices. Because both hemocyanin families contain a similar type-3 binuclear copper site, it is assumed that these proteins both evolved from an ancestral oxygen-binding protein or primitive phenoloxidase (Markl, 2013, van Holde et al., 2001). The binuclear copper site possibly originated from gene duplication and fusion of the Cu-B site. This duplication/fusion event yielded a four- helix bundle, which acquired oxygen-binding ability when O2 accumulated in the atmosphere and a circulating oxygen-transport protein became crucial to utilize the advantages of aerobic metabolism (van Holde et al., 2001). Although it is likely that gene duplication of a Cu-B binding motif created a Cu-A binding motif in the hemocyanin evolution, for the molluscan hemocyanin gene fusion of a Cu-B binding motif with an ancestral Cu-A binding site might also have occurred, in view of the quite divergent amino acid sequences of the Cu-A and Cu-B binding motifs in mollusc hemocyanins. Because of this lack of clear sequence conservation in mollusc hemocyanins, while there is in arthropod hemocyanins, this suggests that mollusc and arthropod hemocyanins are the result of convergent evolution. Both hemocyanin families arose probably independently, with molluscan hemocyanin coming into existence much earlier (about 700-800 million years ago) in evolution than arthropodan hemocyanin (about 550-600 million years ago) (van Holde et al., 2001, Burmester, 2002). The primitive molluscan hemocyanin underwent multiple gene duplications and fusions to yield the current protein, in which 7 to 8 functional units sequentially form a subunit of about ~400 kDa that assembles into a huge decameric or di-decameric cylinder-like quaternary structure (Cuff et al., 1998). For the arthropod hemocyanin precursor the next step in the evolution was the fusion of the α-helical core domain with two other domains, probably involved in cooperative O2 binding via inter- subunit interactions, and the assembly into oligomers of hexamers, with each hexamer

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composed of similar or identical subunits with a Mr of ~75 kDa (van Holde et al., 2001, Decker et al., 2000). The independent evolution implies that both hemocyanins originated from distinct though similar precursors. The dissimilar evolutionary pathway of the hemocyanins manifests itself mainly at the Cu-A site. In the molluscan hemocyanin structures, one of the copper-coordinating histidine ligands at the Cu-A site is provided from a loop. This is distinctly different from arthropodan hemocyanins, where the corresponding His residue is located on the same α-helix as where the second Cu-A- coordinating histidine resides (Fig. 4) (Cuff et al., 1998). A second distinguishing difference is the presence of a negatively charged residue 4 or 7 residues after the third copper-ligating histidine residue (denoted H3) of the Cu-A/Cu-B binding motif. In molluscan hemocyanins, a Glu is present at position H3+7 of the Cu-A binding motif, and an Asp at position H3+4 of the Cu-B binding motif. In arthropodan hemocyanins, the acidic residue is at position H3+7 of both the Cu-A and Cu-B binding motifs (Cuff et al., 1998). Tyrosinases (also called polyphenol oxidases) contain also a type-3 binuclear copper site similar to that of hemocyanins. Since bacterial, fungal, plant, and mammalian (including human) tyrosinases share ~25-30% sequence identity with molluscan hemocyanins, it is generally believed that a primitive molluscan hemocyanin (probably already with some low promiscuous phenoloxidase activity) diverged into present-day molluscan hemocyanin and tyrosinase. Likewise, the primitive arthropodan hemocyanin also diverged into current hemocyanins and prophenoloxidases. Since arthropodan hemocyanins and prophenoloxidases are more similar to each other than molluscan hemocyanins and tyrosinases, the divergence of the latter is more ancient (van Holde et al., 2001). The arthropod-specific prophenoloxidases do not closely resemble their non- arthropod paralogues (Burmester, 2002), indicating a quite different evolutionary history. For instance, the amino acid sequence identity of Agaricus bisporus tyrosinase, of molluscan origin, to Manduca sexta prophenoloxidase is only 13%. On the basis of sequence similarities two distinct phenoloxidase families can be defined, tyrosinases and prophenoloxidases, respectively from molluscan and arthropodan origin, like the two hemocyanin families. A related class of proteins, belonging to the arthropod hemocyanin superfamily, is formed by the hexamerins, which are proteins found in () and which are thought to function as storage proteins, serving as a source of amino acids and energy for protein synthesis during metamorphosis of the larvae (Burmester et al., 1996). Some of them contain a large amount of aromatic amino acids (arylphorins), whereas others have high amounts of methionine (Burmester, 2002). Hexamerins and arthropod hemocyanins share ~30% sequence identity in the core, and likely originate from a common ancestor. However the hexamerins have lost the copper-binding ability because all six copper-ligating histidine residues are mutated (Beintema et al., 1994).

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Probably, their oxygen-binding property became obsolete because insects have a tracheal respiratory system using diffusive O2 transport, and thus have no need for a specific oxygen-transporting protein. Therefore, it has been proposed that hexamerins changed their function to storage after losing the capability of copper-binding (Burmester et al., 1996). However, recently also hemocyanins have been described that could be involved in O2 transport under certain environmental conditions or during some developmental stages (Pick et al., 2009). Likely, the co-existence of copper-less and copper-containing hemocyanin-like proteins may have promoted the emergence of hemocyanin-like storage proteins in insects. Molluscan and arthropod hemocyanins are a clear example of convergent evolution, with two independent gene duplication events, albeit probably starting from the same copper-binding motif. Thereafter, further divergent evolution led to the emergence of tyrosinases, and in the arthropodan family, hexamerins and prophenoloxidases.

23

Chapter 1 ......

24

General introdution ......

Figure 4. Cartoon representation of the crystal structures of (A) Agaricus bisporus tyrosinase (PDB ID 2Y9W (Ismaya et al., 2011)); the His residues, which coordinate the binuclear copper site, are depicted as sticks in the zoomed-in window, (B) Octopus dofleini (mollusc) hemocyanin functional unit (PDB ID 1JS8 (Cuff et al., 1998)), (C) Antheraea pernyi hexamerin-like arylphorin (PDB ID 3GWJ (Ryu et al., 2009)) and (D) Panulirus interruptus (arthropod) hemocyanin (PDB ID 1HC1 (Volbeda et al., 1989). The four α-helices that make up the active site are depicted in green (Cu-A binding motif) and red (Cu-B binding motif). The brown spheres are the Cu-A (left) and Cu-B (right) copper centres. For viewing purposes the lectin-like domain in tyrosinase (A) and the N-terminal domains of (C) and (D) have been omitted.

2.3.3 Other oxygen-transport proteins

A third class of oxygen transport proteins is formed by the hemerythrins. Hemerythrins are responsible for oxygen transport in the marine invertebrate phyla of sipunculids (peanut worms), priapulids (penis worms) and brachiopods (shellfish). Hemerythrin has also a four α-helix bundle core with a pair of iron ions binding O2. The protein occurs in various multimeric states (monomers, trimers and octamers (Meyer et al., 2010)). The iron ions are bound to the protein through five histidine residues as well as through the carboxylate side chains of a glutamate and an aspartate residue (Holmes et al., 1991). The evolutionary history of hemerythrins is characterized by frequent gene losses (Martín-Durán et al., 2013). The examples described in sections 2.2.1, 2.3.2, and 2.3.3 show that oxygen-transport proteins have evolved in several independent ways, making use of different metal ions (Fe2+ or Cu+), and different metal-coordination schemes. From this we conclude that the present-day oxygen-transport proteins are the result of convergent evolution.

2.4 Parallel evolution

Parallel evolution is the appearance of similar functional and structural properties within protein families that evolved in separate lineages from a distant, ancient ancestor. In a remarkable number of cases, parallel evolution has occurred by the repeated acquisition of precisely the same mutations, sometimes in the very same order, which indicates that constraints strongly limit the set of accessible sequences that can produce the selected phenotype (Harms et al., 2013). However, at the DNA and protein sequence level, it is rare to find parallel evolution. Nevertheless, two cases can be noted. One of these cases is exemplified by the evolution of hemoglobin in birds living in the high altitudes of the Andes and other high-mountain areas. Five lineages of birds can be distinguished that independently evolved hemoglobin variants with a higher oxygen affinity than the hemoglobins of low-altitude birds. These hemoglobin variants evolved in parallel in distantly related lineages. Surprisingly, in all variant hemoglobins the

25

Chapter 1 ......

amino acid substitutions are concentrated in the same few regions of the protein (McCracken et al., 2009). Parallel evolution has also been observed in haloalcohol dehalogenases. Haloalcohol dehalogenases catalyze the -independent dehalogenation of vicinal haloalcohols. Crystal structures revealed that they are structurally related to short-chain dehydrogenase/reductase (SDR) enzymes (with up to 30-35% sequence identity), but that they lack the NAD+-co-enzyme binding site (Kavanagh et al., 2008). Two families of haloalcohol dehalogenases (HheA/C and HheB) 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 and diverged via parallel evolution (de Jong et al., 2003). Finally, as described in section 2.3.2, the development of molluscan hemocyanins and tyrosinases on the one hand and arthropodan hemocyanins, prophenoloxidases and hexamerins on the other hand can be considered as independent parallel evolution as well.

2.5 Catalytic promiscuity (poly-reactivity)

Catalytic promiscuity is the ability of an enzyme to accelerate another reaction (or reactions) in addition to its biologically relevant one. Promiscuity of enzymes is essential for the evolution of new enzymatic functions. It has been speculated that superfamily ancestors exhibited broad substrate specificity and could possibly catalyze a range of reactions (Khersonsky et al., 2010). Enzymes often possess other, secondary activities. Under selective pressure, the gene for the existing enzyme may be duplicated, followed by the introduction of a small number of mutations. As a result the secondary activity may become a new main activity (Poelarends et al., 2006). The existence of catalytic promiscuity provides the cell with an effective advantage for the survival of the species and its evolution. A good example of catalytic promiscuity is given by penicillin acylase. Penicillin G acylases (PGAs) are robust industrial catalysts used for the biotransformation of beta- lactams into key intermediates for the production of beta-lactam antibiotics. Their catalytic promiscuity is obvious from the different activities they display, such as trans- esterification (transformation of an ester into another ester through an interchange of the alkoxy moieties), Markovnikov addition (electrophile addition to a double bond of a nucleophile) or Henry reaction (formation of carbon–carbon bond in a reaction of a nucleophilic nitroalkane with an electrophilic aldehyde or ketone) (Grulich et al., 2013). Catalytic promiscuity should not be confused with broad substrate specificity. For instance, P450s and glutathione S- are involved in the detoxification of a wide range of harmful compounds. Being able to detoxify a broad

26

General introdution ...... range of substrates is inherent to their function, and is therefore not to be considered as catalytic promiscuity (Khersonsky et al., 2010, Copley, 2003). The interesting further examples of catalytic promiscuity reviewed in the next section come from various previous and ongoing research projects of the Laboratory of Biophysical Chemistry of the University of Groningen in collaboration with other research groups.

2.5.1 The α/β hydrolase fold superfamily

The α/β hydrolase fold superfamily of enzymes was proposed on the basis of similarities in 3D structures and conservation of the position of the active site residues with respect to the secondary structure elements (Ollis et al., 1992, Nardini et al., 1999a, Carr et al., 2009). The family contains enzymes that, in principle, catalyze a hydrolysis reaction, although also other activities have been observed such as bromoperoxidase activity (Hecht et al., 1994). The canonical fold of this highly divergent enzyme superfamily consists of a highly twisted, 8–11-stranded, mostly parallel -sheet. This sheet is flanked on both sides by α-helices. Additional motifs of variable size, structure and position may decorate the core domain (Marchot et al., 2012). These enzymes have diverged from a common ancestor and preserve the arrangement of the catalytic residues, but not the substrate-binding site. They all have a catalytic triad, the elements of which are borne on loops, which are the best-conserved structural features of the fold (Ollis et al., 1992, Carr et al., 2009).

Figure 5. The canonical α/β hydrolase fold. α-Helices and β-strands are represented by rectangles and arrows, respectively. Locations, where insertions in the canonical fold may occur, are indicated by dashed lines.

Enzymes belonging to this superfamily include serine carboxypeptidases, peroxidases, esterases, lipases (Nardini et al., 2000, Lang et al., 1998, van Pouderoyen et al., 2001,

27

Chapter 1 ......

Tiesinga et al., 2007, Rozeboom et al., 2014), cutinases, thioesterases, dienelactone hydrolases, epoxide hydrolases (Nardini et al., 1999b) and haloalkane dehalogenases (Franken et al., 1991, Verschueren et al., 1993, Ridder et al., 1999). Especially the lipases of the superfamily have been widely applied in biotechnology because of their broad substrate-specificity, and regio- and stereospecificity. (Jaeger et al., 2002). Lipases have also been shown to catalyze other (promiscuous) reactions such as formation of carbon-carbon, carbon heteroatom, and heteroatom-heteroatom bonds through non-hydrolytic reactions, such as aldol condensations and Michael-type additions, and amide hydrolysis by cleavage of a different bond (C-N versus C-O) (Kapoor et al., 2012). Because of their broad repertoire of catalyzed reactions several industrial processes based on lipase catalysis have been established (Jaeger et al., 2002).

2.5.2 Tautomerases

The tautomerase/MIF superfamily members catalyze a keto-enol tautomerisation reaction, promoted by an amino-terminal proline; they all exhibit a two-layered β-α-β- fold structure. The superfamily can be divided into five families, each named after a representative reference enzyme, viz. macrophage migration inhibitory factor (MIF), malonate semialdehyde decarboxylase (MSAD), 4-oxalocrotonate tautomerase (4-OT) (including trans-3-chloroacrylic acid dehalogenase, CaaD), cis-3-chloroacrylic acid dehalogenase (cis-CaaD), and 5-carboxymethyl-2-hydroxymuconate (CHMI) (Baas et al., 2013). Although members of the five families have little overall sequence similarity, the catalytic amino-terminal proline residue is absolutely conserved in all sequences (Poelarends et al., 2008a). Most family members are homotrimers, only CaaD is a heterohexamer with 3 α- and 3 β-subunits, and 4-OT is a hexamer (Poelarends et al., 2008a). Several members of the superfamily display promiscuous activities. For instance, trans- CaaD and cis-CaaD primarily catalyze the hydrolytic dehalogenation of trans-3- chloroacrylate and cis-3-chloroacrylate, respectively, to yield malonate semialdehyde. Both enzymes have catalytic promiscuity in that they also catalyze the tautomerization of phenylenolpyruvate to phenylpyruvate and the hydration of 2-oxopent-3-ynoate (2-OP) to acetopyruvate (Baas et al., 2013). Both enzymes seem to have evolved within a few decades and appear to be products of rapid evolution (Baas et al., 2013). This suggests a very fast ticking molecular clock under the evolutionary pressure of the xenobiotic nematocide 1,3-dichloropropene. Another example of a promiscuous enzyme in the superfamily is Cg10062 from Corynebacterium glutamicum. Cg10062 is a cis-CaaD homologue, which catalyzes the hydrolytic dehalogenation of both cis- and trans-3-chloroacrylate. In addition, the enzyme has hydratase activity, but its major activity is the dehalogenation of cis-3-

28

General introdution ...... chloroacrylate (Baas et al., 2013); the trans-CaaD and hydratase activities can be considered as promiscuous activities. Because of its 34% sequence identity to cis-CaaD, the enzyme has been suggested to be an intermediate in Corynebacterium glutamicum in the evolution to a fully functional cis-isomer-specific dehalogenase such as cis-CaaD (Baas et al., 2013, Poelarends et al., 2008b). The promiscuous activities provide evidence for divergent evolution from a so far unknown common ancestor. An interesting and perhaps unexpected member of the tautomerase superfamily is the macrophage migration inhibitory factor (MIF). The mammalian MIF factor plays an important role in inflammatory and immune-mediated diseases and triggers an acute immune response. Mammalian MIF is also known as a very efficient phenylpyruvate tautomerase and has promiscuous trans-3-CaaD activity (Baas et al., 2013). The MIF homologue from the marine cyanobacterium Prochlorococcus marinus has also tautomerase activity and shares mechanistic and structural similarities with mammalian MIFs (Wasiel et al., 2010). As mammalian MIF was presumably recruited long ago to serve a non-enzymatic function, it is quite remarkable that it still exhibits promiscuous enzymatic activities typical of the other superfamily members (Poelarends et al., 2008a). This suggests that the cyanobacterial MIF and mammalian MIFs are related by divergent evolution. The question arises whether a MIF-like protein from a free-living bacterium also possesses immunostimulatory features similar to those of mammalian MIFs (Wasiel et al., 2010), but we are not aware of any experiments aiming to shed light on this question.

Figure 6. Ribbon diagrams of crystal structures of members of the tautomerase family. (A) Prochlorococcus marinus MIF (PDB ID: 2XCZ (Wasiel et al., 2010), (B) Corynebacterium glutamicum Cg10062 (PDB ID: 3N4G (unpublished), (C) Pseudomonas pavonaceae CaaD (4-OT) (PDB ID: 1S0Y, (de Jong et al., 2004), (D) Pseudomonas pavonaceae MSAD (PDB ID: 2AAG (Almrud et al., 2005), (E) E. coli CHMI (PDB ID: 1OTG (Subramanya et al., 1996).

2.6 Remaining questions

Experimental resurrection of ancestors of universal proteins that have been maintained in nearly all organisms throughout the history of life, together with the sensitivity of single-molecule techniques, can be a powerful tool for understanding the origin and

29

Chapter 1 ......

evolution of life on Earth. Paleoenzymology is the study of prehistoric life through the properties of enzymes. Important pathways such as energy production, sugar degradation, cofactor biosynthesis and amino acid processing are highly conserved from bacteria to humans and were probably already present in the last universal common ancestor. Resurrection of extinct proteins is aimed to obtain information on the effects of historical mutations on functional diversification. For example, in a thioredoxin (Trx) resurrection study sequences predicted by phylogenetic analysis were used for reconstruction of probable variants. It appeared that ancient Trxs use chemical mechanisms of reduction similar to those of modern enzymes but have adapted to the their environment from ancient to modern Earth by adapting their activity to the temperature and pH conditions of present-day Earth (Perez-Jimenez et al., 2011). Additional reconstruction studies with other enzymes may provide additional insights on the evolution of ancient phenotypes and metabolisms. Experimental determination of protein function lags far behind the rate of sequence and structure determination. Nature's strategies for evolving catalytic functions can be deciphered from the information contained in expanding protein sequence databases. However, the functions of many proteins in the protein sequence and structure databases are either uncertain (too divergent to assign function based on homology) or unknown (no homologs), thereby limiting the utility of the databases. Furthermore, these databases are probably far from complete, given the current rate of discovery of new sequences from the metagenome (Kahn, 2011). Nevertheless, the enormous enzyme diversity leaves a huge amount of effort to be done on enzyme functional characterization, to allow further advances in medicine, chemistry, synthetic biology, and industry. The structural information presented in this thesis is a pertinent contribution of how enzymes can evolve. The 3D structures discussed in the thesis provide insights how altered active sites in α-1,4 glucan lyase, endo-xylogalacturonan hydrolase, and pyrroloquinoline-quinone-dependent alcohol dehydrogenase change the product and substrate specificity compared with homologous enzymes. The structures of the esterases reveal how enantioselectivity can arise through divergent evolution. Furthermore, the structure of mannuronan epimerase shows that its catalytic machinery has evolved through convergent evolution.

30

General introdution ......

3 SCOPE OF THIS THESIS

3.1 X-ray crystallography

For understanding the catalytic mechanisms of enzymes, high-resolution structures as determined by X-ray crystallography are a prerequisite. Nowadays, X-ray diffraction techniques have passed the state of pioneering techniques. Although still a highly technical process, it is no longer abstruse and time-consuming (Morin et al., 2013). Yet, there is still nothing routine about determining the complete three-dimensional structure of a protein (Alberts et al., 2002). Early X-ray crystallographers referred to attaining the 3D-coordinates of a protein as “solving” the structure, reflecting their perception of the process as a kind of intricate scientific puzzle of many parts. Today, the preferred terminology is “structure determination”, denoting the more prescribed and rigorous methods typically employed (Morin et al., 2013). By the end of 2013 there were almost 100,000 structures deposited in the PDB (Protein Data Bank) including about 85,000 determined by X-ray crystallography. The number of structures deposited each year in the PDB is growing exponentially since the year 2000, when only 10,696 structures had been deposited. X-ray crystallography is utilized as a tool to study macromolecular structures and to aid in elucidating the mechanism of action of enzymes. However, the structure-function analysis includes more than just crystallization, X-ray crystallography and structure analysis. Proteins have to be produced (cloning, bacterial culture growth, overexpression, mutagenesis). The produced proteins have to be purified by means of chromatography (affinity, cation/anion exchange chromatography, gel filtration, etc.). Activity tests and and inhibition experiments are often done, and conformation analysis is carried out using, for instance, dynamic light scattering and SAXS (small-angle X-ray scattering). All these experiments serve to obtain a protein preparation of the highest quality. X-ray crystallography requires that the protein be subjected to conditions that cause the molecules to aggregate into a large, perfectly ordered crystalline array, a protein crystal. Each protein behaves quite differently in this respect, and protein crystals can be generated only through exhaustive trial-and-error methods that often may take many years to succeed, if they succeed at all (Alberts et al., 2002, Van der Klei et al., 1989, Boys et al., 1989). The first enzyme molecule to be crystallized in 1926 was jack bean urease by the American biochemist J. B. Sumner for which he was awarded the Nobel Prize in chemistry in 1946. It took another 83 years before its crystal structure was finally reported (Balasubramanian et al., 2010). Nevertheless, protein crystallization successes

31

Chapter 1 ......

have multiplied rapidly in recent years owing to the advent of practical, easy-to-use screening kits and the application of laboratory robotics (McPherson et al., 2014). Nowadays, protein crystals are scooped up by a loop, which is made of nylon or plastic and attached to a solid rod. Crystals are pre-soaked in a cryoprotectant, and then flash- cooled with liquid-nitrogen to prevent radiation damage. As X-ray sources the synchrotrons in Grenoble (ESRF) and Hamburg (EMBL outstation at DESY) are used, complemented with the in-house Cu-Kα radiation source, equipped with a MarDTB Goniostat and DIP2030H image plate detector. The major challenges to determine the phase of each reflection in the diffraction data sets of the proteins discussed in this thesis have been overcome either by multiple isomorphous replacement (MIR, soaking crystals in solutions of heavy atoms (see chapter 2)) or by Molecular Replacement (MR, making use of previously determined structures of homologous proteins as phasing models (see chapters, 3, 4, 5 and 6)). Model building and phase refinement is done with freely available programs and as a final step the atomic coordinates of the final models and structure factors are deposited in the PDB.

3.2 Enzymes

The enzymes discussed in this thesis are a heterogeneous group of enzymes, although each protein is of interest by itself because of its unique way of substrate conversion. The six different proteins, each belonging to a different class of enzymes, specifically belong to the classes of (E.C.1 ~9.000 structures in the PDB), hydrolases (E.C.3 ~20.000), (E.C.4 ~ 4.000) and (E.C.5 ~2.500). They are grouped in folds, the α/β hydrolase fold (Chapter 6) and the TIM (β/α)8-barrel (Chapter 3). Other folds discussed are the 8-bladed β-propellers (Chapter 4) and the right-handed β-helix (Chapters 2 and 5).

3.2.1 Evolution of reaction specificity (different products)

3.2.1.1Mannuronan C-5-epimerase (AlgE4)

In Azotobacter vinelandii a family of 7 secreted and calcium-dependent mannuronan C-5 epimerases (AlgE1-7) has been identified (Aachmann et al., 2006). Alginate is a family of linear copolymers of (1-->4)-linked β-D-mannuronic acid (M) and its C-5 epimer α-L- guluronic acid (G). The polymer is first produced as polymannuronic acid and the guluronic acid residues are then introduced at the polymer level as blocks by mannuronan C-5-epimerases. This reaction is catalyzed by mannuronan C-5-epimerase, which abstracts the proton at C-5. The reaction is completed by re-donation of a proton to C5 from the opposite face of the sugar ring.

32

General introdution ......

The smallest of the AlgE proteins, AlgeE4, comprises one A- followed by one R-module and strictly forms alternating MG sequences (MG-blocks). The A-modules alone are catalytically active, but their reaction rates are significantly increased when covalently bound to at least one R-module (Ertesvåg et al., 1999). The structure of the A-module, a right-handed parallel β-helix structure was elucidated by X-ray crystallography (Chapter 2). The catalytic machinery displays a stunning resemblance, presumably through convergent evolution, to the structurally unrelated alginate lyase A1-III from family PL5 that contains an (α/α)6 barrel and alginate lyase ALY-1 from family PL7 containing a β-jelly roll.

3.2.1.2.α-1,4 Glucan lyase (GLase)

The crystal structure of GLase is described in chapter 3 in the native form and in complex with the inhibitors 1-deoxynojirimycin, acarbose, 5-fluoro-β-L-idopyranosyl fluoride (5FβIdoF) and 5-fluoro-α-D-glucopyranosyl fluoride (5FαGlcF). The unique lytic enzyme α-1,4-glucan lyase (EC 4.2.2.13) has been assigned to glycoside hydrolase family

GH31, which consists of mainly hydrolases (EC 3.2.1.x). The enzymes contain a (β/α)8- barrel as core domain with the active site at the C-terminal ends of the barrel strands.

The (β/α)8-barrel, which was found first in triosephosphate isomerase and therefore is also known as TIM barrel, is the most common enzyme fold. (β/α)8-Barrels have high structural homology but their overall sequence identities can be insignificant. Structural homology suggests a common evolutionary origin for the four major classes of GH31, which, however, have diverged to meet the substrate specificity as desired for their environment. GLase represents an example of a way in which enzymatic mechanisms can evolve through subtle changes in active site constitution.

3.2.2 Evolution of substrate specificity

3.2.2.1 Pyrroloquinoline quinone dependent alcohol dehydrogenase (PQQ-ADH)

Pyrroloquinoline quinone dependent alcohol dehydrogenase (PQQ-ADH) belongs to the E.C. 1.1.5 family of oxidoreductases. Its 3D structure elucidation and characterization are described in chapter 4. PQQ-ADH belongs to the PQQ family of β-propeller-fold proteins. β-Propeller proteins adopt folds comprising 4 to 10 repeats of a 4-stranded β-meander called a ‘blade’. They are thought to have largely arisen by the independent amplification of one ancestral blade and not from an ancestral propeller. The differentiation of the first proto-propellers yielded a population of new β-meanders, which could serve as starting points for further amplification. This evolutionary pathway of amplification and diversification is an ongoing process (Chaudhuri et al., 2008).

33

Chapter 1 ......

The phylogenetic relationship of the PQQ-domains of ten structurally characterized PQQ- dependent ADHs with eight 4-stranded β-meanders is shown in Fig. 1. Type II QH-ADHs are structurally closely related to QADHs, from which we infer that they are also evolutionary related. Somewhat more distant from these types are type I Methanol Dehydrogenases. Our results show that PQQ-ADH is the most distant to all mentioned types and could be regarded as an outgroup in this tree. This suggests that PQQ-ADH branched from the universal ancestor before the other enzymes branched from each other. Yet, the number of samples is limited in this particular example and the tree may be altered when new 3D structures become available.

3.2.2.2 Endo-Xylogalacturonan hydrolase (XghA) from Aspergillus tubingensis

XghA belongs to E.C. 3.2.1.-, (no subfamily assigned) and is a member of Glycoside Hydrolase family GH28, which contains polygalacturonases and rhamnogalacturonases. Family GH28 is a set of structurally related enzymes with interesting functional diversity that hydrolyze glycosidic bonds in pectin and exhibit the parallel β-helix topology. The β- helix fold shows extensive stacking of side chains of similar residues in its interior to give aliphatic, aromatic and polar stacks such as the asparagine ladder. This suggests that coils can be added or removed by duplication or deletion of the DNA corresponding to one or more coils and explains how homologous proteins can have different numbers of coils (Jenkins et al., 2001). In most β-helix proteins a surface groove is observed that forms the center of the active site, but the active site is also shaped by additional loops. The structure of Mannuronan C-5 epimerase also contains a β-helix (chapter 2) with a surface groove but the catalytic machinery is completely different. Aspergillus tubingensis and Aspergillus niger are closely related fungi, which both belong to the subgenus Circumdati, section Nigri (Abarca et al., 2004). In A. niger, 20 putative GH28 homologs are found (Sprockett et al., 2011). The evolution of the GH28 gene family within the fungal kingdom is consistent with the birth-and-death model. In the birth- and-death model new genes are created by repeated gene duplication and some duplicate genes are maintained in the genome for a long time but others are deleted or become nonfunctional by deleterious mutations. The initial appearance of GH28 enzymes predates the evolution of fungi, which places the origin of this gene family at more than 1.5 billion years ago. The earliest fungi already possessed much of the functional diversity seen today in GH28, including exo-polygalacturonan-, endo- polygalacturonan-, and endo-rhamnogalacturonan-acting enzymes (Sprockett et al., 2011). In chapter 5 the structure of XghA is described. XghA has probably evolved through gene duplication of an ancestor of GH28 enzymes followed by divergent evolution. Its

34

General introdution ...... substrate-binding cleft can accommodate xylosylated substrates and is much wider than in other polygalacturonases that cannot process xylosylated polygalacturonan.

3.2.3 Evolution of enantioselectivity

3.2.3.1 Esterases (NP and CesB)

Examples of enzymes of which the enantioselectivity has successfully been improved or inverted by rational design are rare. The effects of certain mutations to alter enzyme enantioselectivity are more difficult to predict than mutations focused on altering other catalytic properties. However, directed evolution (evolutive biotechnology or molecular evolution) combined with rational design can be used to improve the enantioselectivity (Bornscheuer et al., 2001, Boettcher et al., 2010). The carboxylesterases naproxen esterase (NP) and CesB perform enantioselective biochemical transformations. NP was characterized as a very efficient enantioselective biocatalyst for the kinetic resolution of esters of the non-steroidal anti-inflammatory drug naproxen, while CesB shows promise for application in the enantioselective production of the chiral alcohol 1,2-O-isopropylidene-sn-glycerol. Although the crystallization of NP from Bacillus subtilis Thai I-8 was already reported in 1993 (van der Laan et al., 1993), it took 20 years to determine its structure, as described in chapter 6. In addition, in this chapter the 3D structure of its homolog CesB from B. subtilis 168 is described. Carboxylesterases NP and CesB belong to class EC 3.1.1.1 (hydrolases) and contain a α/β hydrolase fold with an α-helical cap domain. Many subclasses of the esterases have been described and many structures have determined so far. Both of the determined esterases show structural flexibility necessary to perform their activity. This makes the outcome of just rational design of mutants that should increase enantioselectivity difficult to predict.

Chapter 7 summarizes the main conclusions of the research described in this thesis. Together with the findings from biochemical and functional experiments, the lessons learned from the structures of glucansucrases and fructansucrases provide directions for future research on these intriguing enzymes.

35

Chapter 1 ......

36

Chapter 2

Structural and Mutational Characterization of the Catalytic A-module of the Mannuronan C- 5-epimerase AlgE4 from Azotobacter vinelandii

Henriëtte J. Rozeboom‡1, Tonje M. Bjerkan§1, Kor H. Kalk‡, Helga Ertesvåg§, Synnøve Holtan§, Finn L. Aachmann§, Svein Valla§, and Bauke W. Dijkstra‡

‡Laboratory of Biophysical Chemistry, GBB, University of Groningen, Nijenborgh 4, 9747 AG Groningen, The Netherlands and the §Department of Biotechnology, The Norwegian University of Science and Technology, Sem Sælands vei 6/8, N-7491 Trondheim, Norway

The Journal of Biological Chemistry (2008) 283 23819 –23828

1These authors contributed equally to this article. T.M.B. carried out protein expression, purification, mutagenesis, and enzymatic assays, secondary structure determination, and production and purification of mannuronan oligomers, analysed the data and wrote the paper H.J.R. carried out crystallization experiments, crystallographic data collection and processing, structure determination and refinement, analysed the data and wrote the paper.

......

37

Chapter 2 ......

ABSTRACT

Alginate is a family of linear copolymers of (1→4)-linked β-D-mannuronic acid and its C-5 epimer α-L-guluronic acid. The polymer is first produced as polymannuronic acid and the guluronic acid residues are then introduced at the polymer level by mannuronan C-5- epimerases. The structure of the catalytic A-module of the Azotobacter vinelandii mannuronan C-5-epimerase AlgE4 has been determined by x-ray crystallography at 2.1Å resolution. AlgE4A folds into a right-handed parallel β-helix structure originally found in pectate lyase C and subsequently in several polysaccharide lyases and hydrolases. The β- helix is composed of four parallel β-sheets, comprising 12 complete turns, and has an amphipathic α-helix near the N terminus. The catalytic site is positioned in a positively charged cleft formed by loops extending from the surface encompassing Asp152, an amino acid previously shown to be important for the reaction. Site-directed mutagenesis further implicates Tyr149, His154, and Asp178 as being essential for activity. Tyr149 probably acts as the proton acceptor, whereas His154 is the proton donor in the epimerization reaction.

INTRODUCTION

Alginate is a family of linear copolymers of (1→4)-linked β-D-mannuronic acid (M) and its C-5 epimer α-L-guluronic acid (G) produced by brown algae and some bacteria belonging to the Pseudomonas and Azotobacter genera (Fischer et al., 1955, Drummond et al., 1962, Linker et al., 1964, Gorin et al., 1966). The M- and G-moieties are distributed irregularly as blocks of homopolymeric regions (M- or G-blocks) interspersed with blocks of alternating structure (MG-blocks). The relative amounts and sequence distributions of M and G vary considerably between alginates from different sources, and this variation in sequence and composition imparts different properties to alginates. G- blocks are rather stiff, and form strong, brittle gels with divalent cations such as Ca2+, whereas M-blocks are less rigid and do not form cationic gels. MG-blocks are the most flexible and have been shown to bind Ca2+ and form junction zones (Donati et al., 2005). In Azotobacter vinelandii, alginate constitutes a major part of the coat surrounding cells differentiated into resting stage cysts (Lin et al., 1969, Sadoff, 1975), and alginate negative strains do not encyst (Campos et al., 1996). Nevertheless, the polymer is also produced during vegetative growth, and under such conditions it may play a role in protecting the bacterium from oxidative stress (Sabra et al., 2000). The alginate polymer is produced as polymannuronic acid, and the guluronic acid residues are introduced at the polymer level by the action of mannuronan C-5-

38

Crystal structure of AlgE4A ......

epimerases (Valla et al., 2001). All known alginate-producing bacteria possess a periplasmic C-5-epimerase, AlgG (Rehm et al., 1996), but A. vinelandii also secretes a family of seven extracellular, Ca2+-dependent mannuronan C-5-epimerases, algE1–7 (Ertesvåg et al., 1995, Ertesvåg et al., 1995). These enzymes are highly homologous and consist of one or two catalytic A-modules and one to seven regulatory R-modules, of which at least one is needed for full activity (Ertesvåg et al., 1999). Each of the extracellular A. vinelandii enzymes produces a distinctive M/G-pattern (Ertesvåg et al., 1999); AlgE2 and AlgE5 make relatively short G-blocks, whereas AlgE6 makes longer G- blocks (Svanem et al., 1999, Ramstad et al., 1999). AlgE4 almost exclusively produces alternating MG sequences (Høidal et al., 1999). AlgE4 is the smallest of the extracellular epimerases with a molecular mass of 57.7 kDa (553 amino acids). It contains one A-module, which is active on its own, comprising the 385 N-terminal amino acids, and one R-module of 142 residues, connected by a 7- residue linker. At the C terminus it has a small “S motif” of 19 residues of unknown function (Ertesvåg et al., 1995). Atomic force microscopy studies have revealed that the AlgE4 A-module binds more strongly to the alginate chain than the complete enzyme, indicating that the R-module is involved in the regulation of the enzyme-substrate binding strength (Sletmoen et al., 2004). Nevertheless, the R-module on its own is also able to bind to the alginate chain (Aachmann et al., 2006). AlgE4 has its optimum activity around pH 6.5–7.0 in the presence of 1–3 mM Ca2+ and at temperatures close to 37 °C (Høidal et al., 1999). It is highly sensitive to alkaline pH, being inactive at pH >8. It binds a minimum of 6 sugar residues in the catalytic site, and for short chains the third residue from the non-reducing end is the first to be epimerized (Campa et al., 2004). For longer chains the reaction proceeds toward the reducing end in a non-random, processive manner (Hartmann et al., 2002). On average, the enzyme epimerizes 10 units ((MG)10) in each reaction before leaving the chain (Campa et al., 2004). An alignment of all known mannuronan C-5-epimerase sequences from bacteria and algae showed that they share an YG(F/I)DPH(D/E) motif (residues 149–155 in AlgE4) (Svanem et al., 2001, Nyvall et al., 2003, Douthit et al., 2005). Asp152 of this motif, which is also conserved in some pectate lyases, has been shown to be important for activity in AlgE7 (Svanem et al., 2001). The reaction mechanism of C-5-epimerases has been proposed to be similar to the lyase reaction (Gacesa, 1987), involving three steps. First, the target uronic acid charge is neutralized, after which the proton at C-5 is abstracted. An enolate anion intermediate is formed, which is stabilized by resonance. In the lyase case, the enolate anion leads to β-elimination of the 4-O-glycosidic bond, whereas in the epimerase case β-elimination is prevented by donation of a proton to the opposite face of the sugar ring. Indeed in the bifunctional epimerase/lyase AlgE7 of Asp152 eliminated both activities, suggesting that they occur at the same site (Svanem et al.,

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Chapter 2 ......

2001). Here we report the three-dimensional structure of the AlgE4 A-module, which is responsible for generating MG-alginates, and discuss the location of the active site and a possible catalytic mechanism.

EXPERIMENTAL PROCEDURES

Growth of Bacteria

The bacterial strains and plasmids used in this study are listed in Table 1. Unless stated otherwise, bacteria were maintained at 37 °C in L broth (10 g of tryptone, 5 g of yeast extract, and 5 g of NaCl per liter) or on L agar (L broth supplemented with 15 g of agar per liter). Protein was expressed in 3⨯ L broth (30 g of tryptone, 15 g of yeast extract, and 5 g of NaCl per liter). Media were supplemented with 200 μg/ml ampicillin or 12.5 μg/ml tetracycline when appropriate.

Recombinant DNA Technology

Standard recombinant DNA procedures were done according to Sambrook and Russell (Sambrook et al., 2001), whereas transformations were carried out according to the RbCl transformation protocol (New England BioLabs). Plasmids were isolated using the Wizard® Plus SV minipreps DNA Purification System (Promega). All gene manipulations were done in Escherichia coli DH5α, and the pTB26 plasmid was later transferred to E. coli ER2566 (New England BioLabs). DNA sequencing was performed using the BigDye® Terminator version 1.1 Cycle Sequencing Kit (Applied Biosystems).

Production and Purification of Mannuronan Oligomers

Partial acid hydrolysis of mannuronan was performed essentially as described by Campa et al. (Campa et al., 2004). Briefly, after pre-hydrolysis at pH 5.6, the sample was incubated for 6 h at 95 °C and pH 3.5, and then neutralized with NaOH. The resulting oligomannuronic acid hydrolysis mixture was fractionated on three serially connected columns of Superdex 30 preparative grade (HiLoad 2.6 ⨯ 60 cm, GE Healthcare) at a flow rate of 0.8 ml/min with 0.1 M NH4Ac (pH 6.9) at room temperature. Fractions of 5 ml were collected from three successive column runs and pooled. The pooled samples were stored at 4 °C prior to three cycles of freeze-drying to remove all traces of NH4Ac and to provide solid, purified oligomers of mannuronic acid.

40

Crystal structure of AlgE4A ......

Table 1. Bacterial strains and plasmids

Strains or Relevant characteristicsa Mutagenic oligonucleotideb References Plasmids E. coli DH5 (Sambrook et al., 2001) ER2566 Encodes T7 RNA polymerase New England BioLabs Plasmids pTYB4 IMPACT-CN fusion vector New England containing a C-terminal chitin BioLabs binding Tag pUC7Tc pUC7 containing the tetAR genes (Blatny et al., from RK2, Apr, Tcr 1997) pBL5 Derivative of pTrc99A encoding (Bjerkan et algE4 from Azotobacter vinelandii, al., 2004) Apr pTB22 Derivative of pTYB4 in which a This study NcoI-XmaI fragment of pBL5 containing algE4 was inserted into the corresponding sites of the vector. pTB26 Derivative of pTB22 in which the This study tetAR genes from a pUC7Tc BamHI-fragment was blunt-end ligated into the unique Eco47III site of the vector Tcr. pTB61 algE4 H154F (removing BssSI) CGGCTTCGACCCCttCGAGCAGACC This study pTB62 algE4 Y149F (inserting BspEI) CGAGATGTCCGGaTtCGGCTTC This study pTB63 algE4 Y149H (inserting BspEI) CGAGATGTCCGGacACGGCTTC This study pTB65 algE4 K117R (inserting NotI) AACACCAGCGGCcgcGTCGACGGCTGGTTC This study pTB67 algE4 F122Y (inserting RsaI) CGACGGCTGGTaCAACGGCTATATC This study pTB69 algE4 D173V (removing BglI) CAACGGCCTCGtgGGaTTCGTCGCCGAC This study pTB70 algE4 R195L (inserting BspMI) GCCAACGACCtgCACGGCTTCAAC This study pTB71 algE4 D152E (inserting BstBI) ACGGCTTCGAaCCCCACGAG This study pTB72 algE4 D152N (removing BssSI) CTACGGCTTCaACCCCCAtGAGCAGACC This study pTB73 algE4 H154Y (removing BssSI) CTTCGACCCCtACGAGCAGACC This study pTB74 algE4 R195K (removing BtgI) CGCCAACGACaagCACGGCTTC This study pTB75 algE4 D173N (inserting BsaI) GACAACGGtCTCaACGGCTTCGTC This study pTB76 algE4 D173E (inserting XhoI) CAACGGCCTCGAgGGCTTC This study algE4 D119A (removing SalI, GCAAGGTCGcCGGCTGGTTC This study pTB77 inserting NaeI) pTB78 algE4 P153A (inserting BspHI) CGGCTTCGACgCtCAtGAGCAGACC This study pTB79 algE4 R249A (inserting Eco57I) GACAACGCCgctGAAGGCGTGCTG This study pTB80 algE4 F122V (inserting HpaI) CGACGGCTGGgTtAACGGCTATATC This study pTB82 algE4 D178E (removing SexAI) CGGCTTCGTCGCCGAaTAtCTGGTC This study pTB83 algE4 D178N (inserting NruI) GACGGCTTCGTCGCgaAtTACCTG This study algE4 Q225E (removing BsgI, CCTGGTGGTGgAGCGGGGTCTG This study pTB84 inserting BsrBI) algE4 D152N, D173E; derivative of CAACGGCCTCGAgGGCTTC This study pTB87 pTB72 (inserting XhoI) pTB88 algE4 K255L, Q225E; derivative of CCTGGTGGTGgAGCGGGGTCTG This study pTB81 (inserting BsrBI, removing BsgI) pTB89 algE4 D173E, P153A; derivative of CGGCTTCGACgCtCAtGAGCAGACC This study pTB76 (inserting BspHI)

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Chapter 2 ......

(Table 1, continued)

pTB90 algE4 F122Y, K117T; derivative of CAGCGGCAcGGTaGACGGCTGGTACAAC This study pTB67 (removing SalI) pTB92 algE4 Q225N (removing BsgI) CCTGGTGGTGaAtCGGGGTCTGG This study pTB93 algE4 K255R (inserting BspEI) GTGCTGCTCcgGATGACCAGCGACATCAC This study pTB147 algE4 E155A (removing BssSI) GACCCCCACGcGCAGACCATCAACC This study pTB153 algE4 Y149A (inserting NgoMIV) CGAGATGTCCGGCgcCGGCTTCGACCC This study pTB154 algE4 D152A (inserting BseRI) GGCTACGGCTTCGctCCtCACGAGCAGACC This study pTB155 algE4 H154A (removing BssSI, GGCTACGGCTTCGACCCagctGAGCAGACCATC This study inserting PvuII) pTB156 algE4 D173A (inserting NaeI) CAACGGCCTCGcCGGCTTCGTCGC This study pTB157 algE4 D178A (removing SexAI) CTTCGTCGCCGcaTAtCTGGTCGACAG This study pHE186 algE4 H154R (removing BssSI) GCTTCGACCCaCgCGAGCAGACCATCAAC This study pHE187 algE4 H154K (removing BssSI) CGGCTTCGACCCtaAgGAGCAGACCATC This study pTB185 algE4 H196A (inserting NaeI) CTACGCCAACGACCGCgcCGGCTTCAAC This study pTB186 algE4 H196F (Inserting PvuI) CTACGCCAACGAtCGCttCGGCTTCAAC This study pTB187 algE4 H196Y (Inserting PvrI) CTACGCCAACGAtCGCtACGGCTTCAAC This study pTB188 algE4 K117A (removing SalI) CAACACCAGCGGCgcGGTtGACGGCTGGTTC This study pTB190 algE4 Q156A (removing BssSI) CTTCGACCCCCACGAagcGACCATCAACCTG This study pTB191 algE4 E155A, Q156A (removing CTTCGACCCCCACGcGgcGACCATCAACCTG This study BssSI) pTB192 algE4 Q225A, K255A; derivative of GAAGGCGTGCTGCTCgcGATGACCAGCG This study pTB193 (inserting NruI) pTB193 algE4 Q225A (inserting SmaI) GCCTGGTGGTGgccCGGGGTCTGGAGG This study pTB194 algE4 K255A (removing NruI) GAAGGCGTGCTGCTCgcGATGACCAGCG This study a The abbreviations used are: Apr, ampicillin resistance; Tcr, tetracycline resistance. Plasmids pTB61-194 and pHE186-187 are identical to pBL5 with the exception of the introduced mutation in algE4. b These are the forward primers; the reverse primers were complementary to them. Changed bases are shown in lower case. In pTB85- pTB90 and pTB192 pBL5 was not used as the template.

Protein Expression and Purification

The A-module of AlgE4 was recombinantly expressed from pTB26, a derivative of the IMPACT-CN expression vector pTYB4 (New England BioLabs) with the sequence encoding the first 378 amino acids of AlgE4 inserted (Table 1). AlgE4A was expressed in E. coli ER2566 at 15 °C and purified in a single affinity chromatography step, using the IMPACT-CN Protein Fusion and Purification System (New England BioLabs), as described in the manual.

Secondary Structure Determination

Circular dichroism studies of AlgE4A were carried out with a protein concentration of 5

μM in 100 μM HEPES (pH 6.9), at 293 K, and 5 mM CaCl2 on a JASCO J-175 spectropolarimeter. The resulting spectrum was the average of 12 scans recorded at a rate of 50 nm/min; it was analyzed by the program K2d (Andrade et al., 1993).

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Crystal structure of AlgE4A ......

Crystallization

Crystallization conditions were difficult to find; over 500 different conditions were tested before the first protein crystals appeared. AlgE4A crystals were obtained after two months from hanging drop experiments with 1.6 –1.7 M sodium malonate (pH 7 – 8) as precipitant at room temperature, using drops of 3 μl of protein (11 mg/ml) and 3 μl of reservoir solution (McPherson, 2001). The lozenge-shaped crystals had a typical size of 0.3 ⨯ 0.2 ⨯ 0.1 mm3.

X-ray Data Collection

Before data collection, crystals were soaked for 1 min in a cryoprotectant solution containing 1.7 M malonate and 20% glycerol, directly followed by flash cooling. Data were collected in house at 100 K on a DIP2030H image plate detector (Bruker-AXS, Delft, The Netherlands) using Cu-Kα radiation from a Bruker-AXS FR591 rotating-anode generator equipped with Franks mirrors. Intensity data were processed using DENZO and SCALEPACK (Otwinowski et al., 1997). A highly redundant native data set to 2.1-Å resolution was obtained. The crystals belong to space group P21212 (Table 2). With two monomers of 39.8 kDa in the asymmetric unit, the VM is 3.0 Å3/Da (Matthews, 1968), and the calculated solvent content is 59%. Preparation of heavy atom derivatives was done by conventional soaking experiments (Bluhm et al., 1958) with 1–5 mM of the heavy atom compound. For substrate binding studies crystals were soaked for 18 h in a saturated solution of M4 (an oligomer of four (1→4)-linked -D-mannuronic acid residues) in 50% (w/v) polyethylene glycol 3350, and 10 mM CaCl2 at pH 6.5. The M4 dataset was collected at Bruker-AXS, Delft, using Cu-Kα radiation from a Bruker-AXS Microstar generator with Montel200 multilayer graded optics, and a Proteum-R diffractometer system with a 3-axis goniostat, at a temperature of 100 K.

Structure Determination The structure of AlgE4A was determined by multiple isomorphous replacement with two heavy atom derivatives (Table 2). Heavy atom positions were located with the program SOLVE (Terwilliger, 2003), which was also used for refining the heavy atom parameters and for calculating phases. Density modification and phase extension to 2.1 Å was done with RESOLVE (Terwilliger, 2003), which was also used to automatically build fragments of the main chain. After RESOLVE, the overall figure of merit was 0.45. At this stage, a 2.1 Å electron density map was calculated and visually inspected with the program O (Jones et al., 1991). The map was judged not to be interpretable; no electron density was visible for residues that had not been built by RESOLVE. Moreover, the direction of the main chain could not be deduced and no side chain density was visible. Therefore, in an

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iterative procedure, phases were improved with the ARP/wARP procedure (Morris et al., 2003) combined with model building by hand. When 55% of the main chain had been built in this way, the phases could not be further improved. By assuming that the two heavy atoms from the EMP derivative occupy the same positions in the protein, and by overlaying parts of the already built main chain, the non-crystallographic symmetry operators could be determined. This allowed non-crystallographic symmetry averaging during refinement with CNS (Brunger et al., 1998), and the gradual completion of the model building. The structure refinement converged at 2.1 Å to an Rfree value of 23.5% and a conventional Rfactor of 19.5%. The final model consists of 750 amino acids, residues 2–375 from molecule A and residues 2–377 from molecule B, 520 water molecules, 1 glycerol molecule, and 2 calcium ions. Assuming that the B-values of these calcium ions are similar to those of the neighboring protein atoms the occupancies of the calcium ions are close to 0.5 (Table 3). The stereochemistry of the final structure was evaluated using the program PROCHECK (Laskowski et al., 1993) (Table 3). For the M4 experiment the native structure was used as the starting model. Initial 2Fo - Fc and Fo - Fc maps clearly showed density for an M3 trisaccharide product as well as for calcium and chloride ions. The atomic coordinates of the final models and structure factors have been deposited in the Protein Data Bank (entries 2PYG for the native structure and 2PYH for the M3 structure).

Table 2. Data collection and phasing statistics. Values in parentheses are for the highest resolution shell.

Crystal/derivative Native EMP (C2H5HgPO4) PtCl4 M4 174.0, 121.3, 174.0, 121.2, 174.0, 121.3, 169.6, 121.8, Cell (Å) 44.7 44.6 44.7 45.1 50.0- 2.1 50.0 - 2.8 50.0 - 3.2 Resolution range (Å) 43.6 - 2.7 (2.18 - 2.1) (2.9 - 2.8) (3.31- 3.2) No. of unique refl. 51691(4598) 19544 (1886) 14451 (1368) 26719 Completeness (%) 92.3 (83.6) 81.2 (80.7) 88.5 (87.4) 99.2 (93.8) Overall I/σ (I) 12.4 (1.8) 11.6 (3.5) 11.8 (5.3) 10.7 (1.9) Rsym (%) 9.8 (63.8) 9.6 (30.6) 13.0 (29.7) 10.0 (56.1) No. of sites - 2 1 -

Construction and Characterization of AlgE4 Mutants

Mutants of AlgE4 were constructed by the QuikChangeTM Site-directed Mutagenesis Kit (Stratagene) according to the manufacturer’s instructions using the primers shown in Table 1. AlgE4 mutant proteins (plasmids pTB61–pTB65; pTB67; pTB69–pTB76; pTB78–pTB80; pTB82–pTB84; pTB87; pTB89; pTB92–pTB93; pTB147; pTB153–

44

Crystal structure of AlgE4A ......

Table 3. Refinement statistics. Values in parentheses are for the highest resolution shell.

Crystal/derivative Native M4 Resolution (Å) 50.0-2.1 43.6-2.7 No. of reflections in working set 48,999 25,216 No. of reflections in test set 2,624 1,336 Rwork (%) 19.4 23.3 Rfree (%) 23.5 27.9 Root mean square deviations from ideal Bond length (Å) 0.006 0.007 Bond angles (deg) 1.29 1.30 Ramachandran plot

Non-glycine residues in most favored regions (%) 84.3 Non-glycine residues in additional allowed regions (%) 14.8 Non-glycine residues in generously allowed regions (%) 1.0 Number of non-hydrogen atoms (average B values (Å)2)

total/main Residues in chain/side chain A chain 2–375 1–376 Protein molecule A 2,782 (33.5/33.0/34.0) 2,799 (49.8/49.6/50.0) Residues in chain B 2–377 1–377 Protein molecule B 2,798 (31.0/32.5/31.7) 2,806 (47.5/47.2/47.8) Water 520 (42.0) 61 (36.6) Calcium ions 2 (33.8) 7 (72.5) Glycerol 1 (60.6) Mannuronic acid 3 (107.0) pTB158; pTB185–pTB188; pTB190–pTB194; pHE186–pHE187) and the wild type AlgE4 (plasmid pBL5) were expressed in E. coli DH5α at 37 °C, partially purified by fast protein liquid chromatography on a HiTrap-Q column (GE Healthcare), and their catalytic activities measured in an isotope assay essentially as previously described (Ertesvåg et al., 1999), using two purification buffers and a disruption buffer containing 20 mM MOPS

(pH 6.9) supplemented with 2 mM CaCl2, 2 mM CaCl2, 1 M NaCl, and 4 mM CaCl2, respectively. Protein concentrations were estimated by the Bio-Rad protein assay (Bradford, 1976).

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Chapter 2 ......

RESULTS

Structure of the Mannuronan C-5- epimerase AlgE4 A-module

The crystal structure of the A-module of AlgE4 (AlgE4A) was determined at 2.1-Å resolution by the multiple isomorphous replacement method using two heavy atom derivatives (Table 2). There are two molecules, A and B, in the asymmetric unit. Residues 2–375 from molecule A and residues 2–377 from molecule B could be built into the electron density. Superimposition of the two molecules reveals that they are very similar with 373 Cα atoms overlaying with an root mean square deviation of only 0.20 Å. The AlgE4A molecule folds into a right-handed parallel β-helix structure comprising 12 complete turns with overall dimensions of 67 x 37 x 36 Å (Fig. 1). The number of amino acid residues per turn varies from 20 to 40. The β-helix turns form four parallel β-sheets, named PB1, PB2a, PB2b, and PB3, consistent with the naming of the β-sheets in polygalacturonase II (van Santen et al., 1999). Not being part of the β-helix, residues 307–309 and 316–318 make up a small two-stranded antiparallel β-sheet, whereas residues 19–32 form an amphipathic α-helix that caps the N-terminal end of the β-helix. The C-terminal end of the β-helix is covered by the C-terminal end of the second molecule in the asymmetric unit. This predominantly β-helical structure agrees with circular dichroism studies (Fig. 2), which suggest an α-helix content of 5%, a β-strand content of 47%, and a coil content of 48%, whereas, according to the DSSP algorithm (Kabsch et al., 1983), the crystal structure contains 3.5% α-helix, 46.5% β-strand, and 50% coiled coil. Turns connect the β-strands in adjacent β-sheets. The T1 turns between PB1 and PB2 (a or b) vary in length from 1 to 19 amino acids, whereas the T2 turns between PB2 (a or b) and PB3 are short, consisting of one amino acid residue, except in the first three turns, which comprise 8, 2, and 7 residues, respectively. Between PB3 and PB1, the T3 turns vary from 4 to 12 amino acid residues. The T1 and T3 turns are longer toward the N- and C-terminal parts of the β-helix, respectively, and these extensions form a cleft across the center of the molecule, of which the bottom is made up by PB1. In other β-helix proteins the active site is located in the cleft between the long T1 and T3 loops (van Santen et al., 1999, Jenkins et al., 2004), and also in AlgE4A the catalytically important residue Asp152 (Svanem et al., 2001) is situated in this cleft.

Figure 1. Structure of AlgE4A. A, mannuronan trisaccharide (M3) bound in the active cleft is shown in stick representation. A, the secondary structure elements forming the β-helix fold. The N-terminal β-helix is shown at the top. The 4 sheets forming the helix are colored blue (PB1), yellow (PB2a), green (PB2b), and red (PB3). The T1–T3 turns are also indicated. B, the same structure as in A, seen from the C-terminal end. C, stereo view of the electrostatic potential surface. Positive potential is shown in blue and negative potential in red. The figure was produced using PyMol (DeLano, 2002).

46

Crystal structure of AlgE4A ......

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Chapter 2 ......

A calcium ion with a pentagonal-bipyramidal coordination is bound in the hairpin loop between strands β7 and β8 (Fig. 3), with side chains of Ser91, Thr97, and Asp133 and the main chain carbonyl oxygen atoms of Glu95 and Gly124 coordinating the ion. No other calcium ions were present in the native structure, but after soaking crystals in 10 mM 376 CaCl2 five additional calcium binding sites were identified at the surface close to Glu – Ile357, Glu229, Thr114–Arg111, Asp112–Glu145–Asp169, and Asp308–Thr310. In addition, a chloride ion is bound to Arg90 under these conditions. A glycerol molecule from the cryoprotectant is present in the active site in molecule A, close to His154, Asp178, and Tyr149. Glycerol hydroxyl groups are known to often mimic binding of the oxygen atoms of a saccharide substrate (van Pouderoyen et al., 2003). One cis-peptide is present between Glu155 and Gln156. In proteins in which non-proline cis-peptide bonds have been identified, they often occur at or near functionally important sites and are likely involved in catalysis (Jabs et al., 1999).

Figure 2. Circular dichroism spectrum of 5 μM AlgE4A in 100 μM HEPES (pH 6.9), 20 °C, and 5 mM CaCl2. The spectrum of AlgE4A indicates high β-sheet structure content.

Location of the Active Site

To investigate the interaction of AlgE4A with substrate, crystals were soaked in a solution of M4, a tetrasaccharide of four (1→4)-linked-D-mannuronic acid residues. After overnight soaking electron density was only observed for a trisaccharide (M3), bound in subsites -3, -2, and -1 (subsite nomenclature according to (Yoon et al., 2001), the epimerization reaction takes place in subsite +1). The binding of the trisaccharide does not influence the overall structure of the enzyme; the room mean square deviation of all Cα atoms is only 0.27 Å between native and trisaccharide-bound AlgE4A. The bound trisaccharide adopts a linear conformation and has its non-reducing end in subsite -3 toward the C-terminal end of the protein (Fig. 3; see Table 4 for a summary of the hydrogen bonding interactions between the bound trisaccharide and AlgE4A). A shorter soaking time of 4 h revealed some extra electron density in the +1 subsite, close to

48

Crystal structure of AlgE4A ......

Asp152, a residue previously implicated in catalysis (Svanem et al., 2001), suggesting that during the overnight soaking the substrate had degraded. However, density in the +1 subsite was not clear enough to allow reliable modeling of a tetrasaccharide.

Figure 3. Stereo view of amino acids surrounding the substrate binding site. A bound mannuronan trisaccharide 155 156 (M3) molecule is shown in stick representation. Glu and Gln form a cis-peptide. The image was constructed in PyMOL (DeLano, 2002).

To assess the importance of other amino acids for catalysis and binding of the alginate substrate, numerous mutants of AlgE4 were constructed (Fig. 4), based on its alignment with AlgE1–7 and their essentially inactive homologue AlgY (Svanem et al., 1999). Four residues, Tyr149, Asp152, His154, and Asp178, all located in subsite +1, were found to be absolutely essential for AlgE4 activity. The first three of these amino acids are conserved in all known mannuronan C-5-epimerases, whereas Asp178 seems to be conserved in the A. vinelandii enzymes only. Tyr149, His154, and Asp178 point with their side chains into the +1 subsite (Fig. 3), and are likely to interact with a sugar bound at this subsite. Even conservative replacements of these three residues result in essentially inactive enzymes, strongly suggesting a role in catalysis. Replacement of Asp152 by a Glu makes the enzyme practically inactive and only 1% of the activity remains when it is replaced by an Asn (Fig. 4). Thus, both the charge and the size of this residue are important.

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In the active site, Pro153 and the cis-peptide between Glu155 and Gln156 appear as important elements for providing an exact geometry around the catalytic site. Indeed replacement of Pro153 with Ala resulted in a 10-fold drop in activity. An even more dramatic effect was seen when replacing Glu155 with Ala, leading to a near 1000-fold drop in activity. This residue is unlikely to interact directly with the substrate but the substitution probably leads to the disruption of the cis peptide. In contrast, the replacement of Gln156 with Ala leads to a 10-fold activity drop, whereas replacements by other amino acids (results not shown) result in no detectable activity changes, implying that Gln156 is neither essential for the cis peptide nor for the catalysis, however, in combination with E155A, Q156A leads to an essentially inactive enzyme.

Table 4. Interactions between bound sugar atoms and AlgE4A Sugar atom Protein atom Distance (Å)

M-3 O-5 Lys255 NZ 3.7

M-3 O-6A Lys255 NZ 4.0

M-3 O-6B Lys255 NZ 3.2

M-2 O-2 Gln225 OE1 2.5

M-2 O-3 Lys255 NZ 3.8

M-2 O-5 Gln225 NE2 3.2

195 M-2 O-6A Arg NH2 3.5

His196 NE2 2.9

Gln225 NE2 3.7

M-2 O-6B Arg195 NH1 4.0

M-1 O-1 Tyr149 OH 3.0

M-1 O-2 Tyr149 OH 2.5

M-1 O-3 Asn199 ND2 3.0

195 M-1 O-3 Arg NH2 2.6

M-1 O-6B Leu228 N 2.9

Residues Involved in Substrate Binding

Exchange of Arg195 with Lys reduced the activity of AlgE4 30-fold, whereas its replacement by Leu made the enzyme essentially inactive (0.2% activity) (Fig. 4). Arg195 binds the alginate chain in subsite -1 (Table 4). Its side chain orientation is stabilized by hydrogen bonding interactions with the carboxylate group of Asp173 (2.8 and 3.1 Å). The Asp173 side chain has additional hydrogen bonds with the peptide nitrogen atom of His196 (3.2 Å), and with the N-δ2 atom of Asn199 (3.0 Å). Substitution of Asp173 with the somewhat larger Glu apparently leaves these interactions mostly intact, because this

50

Crystal structure of AlgE4A ......

mutant protein displays only a slight drop in activity, and has limited additive effect in combination with D152N or P153A. In contrast, the D173N mutant has lost most of its activity, presumably due to an impaired interaction with Arg195. Thus Asp173 appears to be important for maintaining a productive conformation of Arg195 for substrate binding, consistent with the activity data for D173A, D173V, R195L, and the corresponding double mutant (Fig. 4). Indeed, the Asp173–Arg195 pair is absolutely conserved in A. vinelandii AlgE1–7. Substitutions of His196 by Ala, or Tyr, reduced the AlgE4 activity 20 – 30-fold, whereas its replacement by Phe resulted in a 100-fold activity drop. The N-ε2 atom of His196 interacts with the M-6OA in subsite -2, implying an important role of 196 His in substrate binding.

Figure 4. Characterization of AlgE4 mutants. Relative activities are shown compared with the wild type activity (100%). Enzymes showing less than 0.05% activity are considered inactive. Only enzymes with decreased activities to less than 60% are considered as being significantly different from the native enzyme.

Gln225 interacts with the alginate chain in subsite -2. Exchange of this residue with Ala, Asn, or Glu reduced the activity about 10–20-fold, demonstrating its importance for substrate binding. The replacement of Lys255 by an Arg decreased the activity only slightly, but the activity decreased considerably when Lys255 was exchanged for an Ala. Lys255 is located near subsite -3 and may interact, either directly or via a water molecule, with the carboxylate of the mannuronic acid residue at subsite -3 (Table 4). The

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importance of these residues for substrate binding is also evident from the nearly total loss of activity for the double mutant Q225A, K255A. Mutants replacing Arg249 in subsite -4, on the other hand, keep most of the activity, implying that this residue is not important for substrate binding. Replacement of Lys117 in the proximity of subsites +1 and +2 into an Arg or Ala reduced the AlgE4 activity 4–6-fold. When it was changed to Glu (the amino acid found in AlgE7) the activity dropped to 3%. The positive charge is thus important at this site. Finally, Phe122 is located close to subsite +2 and probably interacts with the substrate in this site. Replacing Phe122 with Tyr resulted in only a minor drop in activity, whereas the replacement with Val led to a virtually inactive enzyme. The absence of structural data on the substrate binding mode in subsite +2 unfortunately precludes an explanation of these data.

DISCUSSION

AlgE4 is the first structurally characterized β-helix protein that catalyzes a C-5- epimerization reaction. Most β-helix proteins are hydrolases and lyases involved in the degradation of acidic polysaccharides. A Dali structural alignment search (Holm et al., 1993) revealed nine β-helix polysaccharide hydrolases and lyases with three- dimensional structures highly similar to AlgE4A (with Z-scores over 20). However, these proteins do not share any sequence homology that could suggest a common evolutionary origin, and their proposed catalytic residues are not conserved in AlgE4A. In view of the proposed similarities in the catalytic mechanisms of alginate lyases and mannuronan C-5-epimerases, a comparison with the active sites of alginate lyases was made. So far, five alginate lyase crystal structures are available from 3 different polysaccharide lyase (PL) families (see Ref. (Coutinho et al., 1999)). The alginate lyase A1-III from Sphingomonas species A1 (A1-III) is a PL-5 enzyme. Its structure (PDB 1HV6 (Yoon et al., 1999) is predominantly α-helical with a deep tunnel-like cleft, in which a mannuronic acid trimer can bind in an extended conformation (Yoon et al., 2001). The other four crystal structures are the PL-7 alginate lyases PA1167 from Pseudomonas aeruginosa (PDB 1VAV (Yamasaki et al., 2004)), ALY-1 from Corynebacterium sp. (PDB 1UAI (Osawa et al., 2005)), A1-II’ from Sphingomonas sp. A1 (PDB 2CWS (Yamasaki et al., 2005)), and a PL-18 alginate lyase from Alteromonas sp. 272 (PDB 1J1T). Although these enzymes consist, like AlgE4A, almost entirely of β-strands, their structures are quite different from that of AlgE4A. Instead of the parallel β-helix architecture of AlgE4 they display a jellyroll β-sandwich topology with antiparallel β-strands. Osawa et al. (Osawa et al., 2005) noted a striking agreement in spatial arrangement of four amino acid residues in the center of the substrate binding clefts of ALY-1 (PL-7) and A1-III (PL-5). These residues were Gln/Asn, His, Arg, and Tyr. They were proposed to be the catalytic residues, with His functioning as the base in the reaction mechanism. The

52

Crystal structure of AlgE4A ......

same spatial arrangement is also found in the other three lyases, and mutational studies of A1-II’ have indicated that the Tyr residue is particularly crucial for the catalytic reaction (Yamasaki et al., 2005). AlgE4A displays the same spatial arrangement of Asp 152, His154, Lys117, and Tyr149 in its active site (Fig. 5), and the mannuronic trisaccharide binds in a very similar orientation and position to that of the bound trisaccharide in A1-III. Although four active site amino acid residues seem to be positioned similarly, other important residues (particularly Asp178) and the environment at subsites -2 and -3 are very different.

Figure 5, Superimposition of the active site residues of ALY-1 in green (His119, Gln117, Arg72, and Tyr195), A1-III (His192, Asn191, Arg239, Tyr246, and trisaccharide) in lilac, and AlgE4A (His154, Asp152, Lys117, and Tyr149 and trisaccharide) in CPK colors. The image was constructed in PyMOL (DeLano, 2002).

The catalytic mechanism of mannuronan C-5-epimerases has previously been proposed to proceed in three steps: the abstraction of H-5, an inversion of the hexopyranose ring, and the donation of a proton to the opposite side of the pyranose ring (Gacesa, 1987). Tyr149, Asp152, His154, and Asp178 are all positioned in the vicinity of the catalytic subsite (+1); moreover the replacement of these residues with other amino acids generally resulted in essentially inactive enzymes. To analyze the possible role of these residues in 4 catalysis, a mannuronic acid residue in the C1 chair conformation was modeled into subsite +1, by extending the experimentally observed bound M3 trisaccharide chain with an extra M-unit (Fig. 6). The constraints imposed by the active site architecture resulted in only a limited number of possible positions for the +1 mannuronic acid residue without causing steric clashes. The allowed conformations are all very similar, and in all of them the +1 residue is oriented with its C-5 hydrogen atom pointing toward the Tyr149 hydroxyl group. Tyr149 is thus in a position fully compatible with a role as a general base (AA-2 in the model of Gacesa (Gacesa, 1987)), that abstracts the C-5-bound proton from

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the +1 M-residue. At the other face of the +1 sugar His154 is located, in an excellent position to act as a general acid (AA-3 (Gacesa, 1987)) that donates a proton to the C-5 1 atom of the +1 sugar, flipped into the C4 chair conformation (Fig. 7).

Figure 6. 2Fo - Fc electron density of the native active site residues generated from a simulated-annealing composite omit map contoured at 1.5 σ. The M residue in subsite -1 from the M3 structure is superposed on the native structure, the M residue in the +1 site has been obtained by computer modeling. The figure was produced using PyMol (DeLano, 2002).

The Tyr149 hydroxyl group is at hydrogen bonding distance to the Arg195 side chain (held in position by Asp173), which provides a proton relay path to the solvent. This proposed role of Tyr149 resembles the proton abstraction step in chondroitin AC lyase, which also uses a tyrosine residue to abstract a proton, in this case from the C-5 atom of a glucuronic acid (Huang et al., 2001, Rye et al., 2006). In addition, the reaction catalyzed by this latter enzyme proceeds via a similar putative enolic transition state as in the mannuronan C-5-epimerases and alginate lyases (Yoon et al., 2001, Yamasaki et al., 2005). AlgE4 differs from the chondroitin AC lyase, though, in that the glycosidic bond between the -1 and +1 sugars remains intact, and Tyr149 does not donate its proton to the glycosidic oxygen. The essential role of His154 as a proton donor in M → G catalysis explains the lack of activity of AlgE4 at pH > 8 (Høidal et al., 1999), at which pH the His154 side chain presumably is deprotonated.

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Crystal structure of AlgE4A ......

Figure 7. Proposed catalytic mechanism of AlgE4. A and B, the alginate polymer enters the catalytic site. B and C, the carboxylate moiety of the mannuronic acid in subsite +1 is protonated, enabling it to form a hydrogen bond with Asp152 (and/or 178), which stabilizes the substrate-enzyme complex. C, upon deprotonation of Tyr149 (via Arg195) the alkoxide ion group extracts H-5 from the re-face of the mannuronic acid in subsite +1. C and D, a double bond is formed, which makes the conformation of the +1 mannuronic acid partially planar. D, the protonated His154 performs a nucleophilic attack on the C-5 atom of the +1 sugar from the si-face with the concomitant flip of 1 the +1 sugar ring into the C4 chair conformation of guluronic acid. D and E, the carboxylic acid moiety on sugar +1 is deprotonated. E and F, the epimerized sugar leaves the active site and His154 is protonated again. F, the epimerase is ready to perform a new reaction.

An essential feature of the binding mode of the +1 mannuronic acid is that its carboxylic acid moiety points into the interior of the enzyme, and is at hydrogen bonding distance from the carboxylic acid side chains of Asp152 and Asp178. It has been observed that for the activity of pectate/pectin lyases, which proceeds through a similar transition state as the mannuronan C-5 epimerases (Gacesa, 1987), an uncharged or methylated sugar carboxylate significantly enhances reactivity. The close interactions of the +1 carboxylic acid and the presumably negatively charged Asp152 (and/or Asp178) side chain ensures that the sugar carboxylate will be protonated (AA-1 substrate stabilization (Gacesa, 1987)), and thus will be in an uncharged state promoting reactivity. An intriguing question is how AlgE4 prevents the lyase side reaction. In principle, Tyr149 could donate the abstracted proton of the glycosidic oxygen of the bond between the -1

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and +1 uronic acid residue, similar to, e.g. chondroitin AC lyase. We propose that the interaction between Tyr149 and Arg195 may provide an effective pathway to remove the proton from the active site, thus reducing the lyase activity. In addition, the hydrogen bonding interaction between the O-3 hydroxyl group of the +1 sugar and the ring O-5 atom of the -1 sugar residue may help to prevent the two sugars moving apart, which will also diminish the lyase reaction. Although calcium is needed for the activity of AlgE4, no Ca2+ ion is found near the active site of the enzyme, neither in the native state nor after calcium soaking. However, the Ca2+ ion that was identified in the loop between strands 7 and 8 could very well explain the calcium dependence. In the absence of Ca2+ at this position, the loop would probably adopt a different conformation and the active site cleft could be disrupted. Various positively charged protein side chains (Arg90, Lys84, Lys117, Arg195, His196, and Lys255) were identified that could stabilize the negative charge of the carboxylate side chains of the substrate. Of these residues, Arg195 and His196 bind the carboxylate side chain of the sugar residue at subsite -2, whereas Lys255 binds the one at subsite -3. Arg90 binds a chloride ion suggesting that it is also capable of binding the carboxylate side chain of a sugar residue. All these positively charged residues are positioned on one side of the active site cleft. The AlgE4-R-module also has a positively charged cleft on the surface of its β-helix (Aachmann et al., 2006). Connecting the C-terminal residue of the catalytic A-module to the N terminus of the R-module by a peptide bond would create one long β-helix protein, in which a continuous long substrate binding groove runs over the surface of the full-length AlgE4 epimerase (Aachmann et al., 2006), thus rationalizing the contribution of the R-module to sub-strate binding. AlgE4 is a processive enzyme, which binds a minimum of six residues in the active site (Campa et al., 2004). The structure of the A-module shows that the polymer, which has to slide through the active site, is enclosed by two “clamps.” The first clamp is between Gly53 and the Ca2+ binding loop with Tyr93 at the tip, and Arg90 at the bottom. The second clamp is between Arg195 and a loop with Leu228 at the tip, in the -2 subsite. These two clamps span a distance of about 20 Å, which can accommodate 5 sugar residues in a stretched conformation. The +1 site is at about 10 Å from either clamp. If the polymer chain has to slide through these positions it is also clear that, because of steric hindrance, O-2 and O-3 acetylated polymers cannot be epimerized, which explains why AlgE4A has no activity on such substrates. The importance of the second clamp is supported by hybrid enzyme studies in which residues 215 to 262 from AlgE4A, a processive enzyme, were transferred to AlgE2, a non-processiveenzyme,con-verting it into a presumably processive enzyme (Bjerkan et al., 2004). It is still unclear how enzymes with the same catalytic residues distinguish between the formation of MG/GG-blocks and cleavage of the alginate chain. This reaction specificity must come from the environment of the active site, limiting which molecules effectively

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Crystal structure of AlgE4A ......

bind and also which reaction will take place. By comparing AlgE4 to the highly G-block forming enzyme AlgE6 or the bifunctional AlgE7 it is apparent that only subtle differences result in very different activities. Now that the structure of AlgE4 is known, it is possible to form and test hypotheses as to which amino acids are responsible for the various properties. The answers to these questions will also create a deeper understanding of the reaction mechanism of these enzymes. Given the striking similarities of the active sites of the studied epimerases and lyases, the results presented here may also help in understanding substrate binding, and hence the substrate specificities of the different enzymes involved in alginate cleavage or epimerization.

ACKNOWLEDGEMENTS

We thank Drs. A Coetzee and B. Schierbeek for collection of the M4 data at Bruker-AXS and T. Pijning for critical reading of the manuscript.

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Chapter 3

Crystal structure of α-1,4-glucan lyase, a unique glycoside hydrolase family member with a novel catalytic mechanism

Henriëtte J. Rozeboom*, Shukun Yu†, Susan Madrid†, Kor H. Kalk*, Ran Zhang§ and Bauke W. Dijkstra*

*Laboratory of Biophysical Chemistry, Groningen Biomolecular Sciences and Biotechnology Institute, Nijenborgh 7, 9747 AG Groningen, University of Groningen, The Netherlands. †Danisco Innovation, Danisco A/S, DK-1001 Copenhagen, Denmark. §Department of Chemistry, University of British Columbia, 2036 Main Mall, Vancouver, B.C., V6T 1Z1, Canada. Present addresses: S. Yu, Enzyme R&D, Danisco A/S, Braband, Aarhus, Denmark; S. Madrid, Danisco US Incorporation, Palo Alto, CA, USA

Journal of Biological Chemistry, 2013, 37, 26764–26774

......

α-1,4-Glucan lyase (GLase) is a glycoside hydrolase family member that degrades starch via an elimination reaction. Crystal structures of GLase with covalently bound inhibitors show that the catalytic nucleophile can abstract the proton. The nucleophile has a dual function, acting as nucleophile and base. A single substitution converts a glycoside hydrolase into a lyase. This study presents the structure of GLase and reveals a novel role for the nucleophile.

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ABSTRACT

α-1,4-Glucan lyase (EC 4.2.2.13) from the red seaweed Gracilariopsis lemaneiformis cleaves α-1,4-glucosidic linkages in glycogen, starch and malto-oligosaccharides, yielding the keto-monosaccharide 1,5-anhydro-D-fructose. The enzyme belongs to glycoside hydrolase family 31 (GH31), but degrades starch via an elimination reaction instead of hydrolysis. The crystal structure shows that the enzyme, like GH31 hydrolases, contains a (β/α)8-barrel catalytic domain with B and B’ subdomains, an N-terminal domain N, and the C-terminal domains C and D. The N-terminal domain N of the lyase was found to bind a trisaccharide. Complexes of the enzyme with acarbose and 1-deoxynojirimycin, and two different covalent glycosyl-enzyme intermediates obtained with fluorinated sugar analogues show that, like GH31 hydrolases, the aspartic acid residues Asp553 and Asp665 are the catalytic nucleophile and acid, respectively. However, as a unique feature, the catalytic nucleophile is in a position to act also as a base that abstracts a proton from the C2 carbon atom of the covalently bound subsite -1 glucosyl residue, thus explaining the enzyme’s unique lyase activity. One Glu to Val mutation in the active site of the homologous α-glucosidase from Sulfolobus solfataricus resulted in a shift from hydrolytic to lyase activity, demonstrating that a subtle amino acid difference can promote lyase activity in a GH31 hydrolase.

INTRODUCTION

Glycoside hydrolases (GHs) are important biocatalysts that occur in nearly all forms of life, where they catalyze pivotal biological reactions including the hydrolytic degradation of oligo- and polysaccharides (Henrissat et al., 2008). On the basis of their primary structure and conserved sequence motifs, they have been grouped into > 130 glycoside hydrolase families (Cantarel et al., 2009). However, also nonhydrolytic enzymes, such as lyases, also exist and cleave glycosidic bonds. Polysaccharide lyases (EC 4.2.2.-) cleave the O-C4' glycosidic bond by abstraction of the C5’ proton, followed by introduction of an alkene functionality between the C4’ and C5’ atoms (Lombard et al., 2010). In contrast, α- 1,4-glucan lyases (GLases; EC 4.2.2.13) cleave the glycosidic C1-O bond by abstraction of the C2 proton and formation of the enol form of anhydrofructose (Fig. 1) (Yu et al., 1999b). Based on sequence similarity and the cleavage of C1-O glycosidic bonds, GLases have been grouped as special members of glycoside hydrolase family 31 (GH31), instead of as members of a separate polysaccharide lyase family (Yu et al., 1999a). They form GH31 subgroup 2, whereas subgroup 1 contains all known α-glucosidases, including such important enzymes as human lysosomal α-glucosidase, endoplasmic reticulum

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Crystal structure of α-1,4 glucan lyase ......

glucosidase II, and the digestive enzymes maltase-glucoamylase (MGAM) and sucrase- isomaltase. Subgroups 3 and 4 contain archaeal and bacterial α-xylosidases, respectively (Ernst et al., 2006). Sequence alignment shows that GLases have 18-25% identity to the GH31 subgroup 1 α-glucosidases, and less identity to the other subgroups. This alignment further revealed the existence of several signature sequences of subgroups 1 and 2 which include the motifs WxDM and WxGDN (Yu et al., 1999a) The D in motif WxDM was shown to represent the catalytic nucleophile (Lee et al., 2002), while the D in WxGDN functions as the acid/base catalyst (Lovering et al., 2005). Mechanistic studies, based on kinetic isotope effects, suggested that the reaction catalyzed by GLases proceeds via two steps. In the first step a covalent GLase-substrate intermediate (the glycosylated GLase) is formed, which is similar to the first step of the double displacement reaction catalyzed by GH31 retaining glycoside hydrolases. For the second step, a syn elimination of the GLase-substrate intermediate has been proposed, yielding an unsaturated sugar, 1,5-anhydrofructose, and the free enzyme GLase. Both the first and second reaction steps are supposed to proceed via oxocarbenium ion-like transition states (Lee et al., 2002, Lee et al., 2003, Lee et al., 2005). However, which residue abstracts the proton from the C2 atom of the sugar ring in the elimination reaction has remained unclear so far (Lee et al., 2005). GLases are exo-enzymes that act from the nonreducing ends of the substrate. They degrade both malto-oligosaccharides and starch, suggesting that their active site comprises several glucosyl-binding subsites (Yu et al., 1999a, Yoshinaga et al., 1999). Moreover, GLases and glycoside hydrolases share certain competitive inhibitors like 1- deoxynojirimycin and acarbose (Yu et al., 1999a), with Ki values of 0.130 μM (Lee et al., 2003) and 0.002 μM (Lee et al., 2002), respectively. 1,5-Difluoroglycosides are also inhibitors of GLase (Lee et al., 2003). Such sugars are suitably activated at the anomeric carbon with an excellent leaving group. The C5-fluoro group increases the energy of the transition state, thus slowing down both reaction steps catalyzed by the enzyme and therefore these fluorosugars can be used to label the catalytic nucleophile (McCarter et al., 1996). It appeared that epimers of the parent fluorosugars inverted at C5 are more effective trapping reagents for GH38 α-mannosidase (Numao et al., 2003) and GH31 α- xylosidases (Lovering et al., 2005, Larsbrink et al., 2011). Indeed, also for GLase, 5- fluoro-β-L-idopyranosyl fluoride (5FβIdoF) with a Ki of 28.3 ± 2.1 mM (Lee et al., 2002, Lee et al., 2003) is more effective than 5-fluoro-α-D-glucopyranosyl fluoride (5FαGlcF),with an apparent dissociation constant Ki of 6.5 mM (Lee et al., 2003). Here we report the x-ray structure of GLase from Gracilariopsis lemaneiformis, a lyase belonging to GH31, in its native form and in complexes with acarbose, 1- deoxynojirimycin, 5FβIdoF and 5FαGlcF. We show that the catalytic nucleophile Asp553 functions as a nucleophile in the first step of the reaction forming the covalent enzyme- glycosyl intermediate but acts as a catalytic base in the second step, abstracting a proton

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from the C2 carbon atom of the covalently bound subsite -1 glucosyl residue. These results resolve the long time ambiguity on the nature of the proton abstracting residue (Lee et al., 2003, Lee et al., 2005). Site-directed mutagenesis and enzymatic characterization of the archaeal α-glucosidase homologue MalA show that subtle changes are sufficient to confer lyase activity to a hydrolase.

Figure 1. A comparison of the three mechanisms of glycosidic bond breakage, exemplified, respectively, by glucoside hydrolases and two polysaccharide lyases. For anhydrofructose the enol tautomer is shown, which is in equilibrium with the keto form as a result of enol-keto tautomerization (Yu, 2008).

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Crystal structure of α-1,4 glucan lyase ......

MATERIALS AND METHODS

Expression and Purification of α-1,4-Glucan Lyase

The α-1,4-glucan lyase (GLase) gene was cloned from a DNA library using genomic DNA isolated from the red seaweed Gracilariopsis lemaneiformis (Bojsen et al., 1999a) Expression cassettes were constructed and intracellular expression of the GLase gene in the yeast Hansenula polymorpha was achieved as described previously (Cook et al., 2003). Four constructs were made representing full-length, N-terminally truncated, C- terminally truncated and both N-and C-terminally truncated glucan lyases (Fig. 2). Purification of the recombinant glucan lyases was performed by anion exchange chromatography using Q-Sepharose FF with 20 mM Bis-Tris pH 6.7 as Buffer A, and Buffer A containing 1.0 M NaCl as Buffer B (Yu et al., 1995, Yu et al., 1997). Further purification was achieved by a second anion exchange chromatography step using a MonoQ column and elution with a gradient of 0.28 - 0.32 M NaCl in 20 mM Bis-Tris, pH 6.7. Only the first fractions of the GLase peak from subsequent runs (with 1 mg protein per run) were pooled and concentrated to 9.7 – 17.0 mg protein ml-1. Characterization of GLase with respect to its substrate specificity, N-terminal sequencing and mass spectrometry analysis (MALDI-TOF) was as described before (Yu et al., 1995).

Figure 2. Construction of 4 GLase expression cassettes. FL-gene, coding for the full-length glucan lyase gene (1- 1038), CC-gene coding for the N-terminal truncated lyase (224-1038), NC-gene coding for the C-terminal truncated lyase (1-938), and C-gene coding for both N-and C-terminal truncated lyase (224-938). These constructs were used to transform Hansenula polymorpha by electroporation

Crystallization and X-ray data collection

GLase crystals were obtained from hanging-drop experiments with 18-21% polyethylene glycol 8000 in 100 mM sodium acetate buffer (pH 5.0–5.2) as precipitant, using drops of 3 μl protein solution (12.9 mg ml-1 in 20 mM Bis-Tris buffer, pH 6.7, 300 mM NaCl) and 3

μl of reservoir solution. Crystals grew in two different space groups (P1 and P21); both forms were used in the structure determinations (Table 1).

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Before data collection, crystals were soaked for 15 s in a cryoprotectant solution, consisting of reservoir solution supplemented with 18% glycerol, directly followed by flash freezing. Native GLase data were collected at 100 K from a crystal belonging to space group P1 on beam line BW7A (DESY/EMBL, Hamburg, Germany). For inhibitor binding studies, crystals of space group P21 were soaked in a solution containing 15 mM acarbose (Serva Electrophoresis GmbH, Germany) or 125 mM 1-deoxynojirimycin (Sigma-Aldrich) for a few minutes. Data were collected in house at 110 K with a DIP2030H image plate detector using Cu-Kα radiation from a Bruker-AXS FR591 rotating-anode generator equipped with Franks mirrors for the acarbose dataset or with Osmic mirrors for the 1-deoxynojirimycin dataset. Intensity data were processed using DENZO and SCALEPACK (Otwinowski et al., 1997). For the 5-fluorosugar binding studies, crystals of space group P1 were soaked in solution containing 20 mM 5FβIdoF for 30 minutes or 40 mM 5FαGlcF for 40 minutes. 5FαGlcF and 5FβIdoF were synthesized as described previously (McCarter et al., 1996) Data were collected on beam line ID14-4 (ESRF, Grenoble, France) for 5FβIdoF and on beam line ID23-2 (ESRF) for 5FαGlcF. Intensity data were processed using iMosflm (Battye et al., 2011) and scaled using SCALA from the CCP4 suite (Winn et al., 2011) for 5FβIdoF, and with XDS (Kabsch, 2010) for 5FαGlcF. A summary of data collection statistics is given in Table 1.

Structure Determination and Refinement The structure of native GLase was solved by molecular replacement with PHASER (McCoy et al., 2007) using a model of the MalA structure (PDB code 2G3M (Ernst et al., 2006)) made by the FFAS server (Jaroszewski, L., Rychlewski, L., Li, Z., Li, W. & Godzik,A., 2005) as a search model. In an iterative procedure, the phases were improved with NCSREF (Winn et al., 2011, Murshudov et al., 2011) and the ARP/wARP procedure (Langer et al., 2008), combined with manual model improvement with COOT (Emsley et al., 2010). Refinement with REFMAC5 (Murshudov et al., 2011) with TLS rigid body refinement (Painter et al., 2006) as last step, resulted in a final model comprising 4 protein molecules with 1025 residues each, 15 glycerol molecules, 9 acetate molecules, 2 chloride ions and 3766 water molecules. Further details of the refinement are listed in Table 1. The electron density of the first 2 residues from all 4 GLase molecules is not visible.

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Crystal structure of α-1,4 glucan lyase ......

Table 1 Data collection and structure refinement statistics.*Values in parentheses are for the highest-resolution shell.

wt GLase Acarbose soak Deoxynojirimycin 5FIdoF soak 5FGlcF soak soak Data collection

Spacegroup P1 P21 P21 P1 P1 Unit cell a, b, c (Å), 91.7, 96.6, 134.9, 91.9, 134.7, 91.8, 92.1, 97.3, 91.6, 97.0, α, β, γ (o) 134.5 193.5 193.2 136.3 135.7 80.5, 83.3, 85.3 90.0, 99.4, 90.0 90.0, 99.3, 90.0 80.3, 83.3, 85.2 80.4, 83.1, 85.2 Resolution (Å) 2.06 2.60 2.35 1.90 2.10 No. of observations 3213515 1914405 1569071 663997 477958 No. of unique refls. 258096 322607 447794 340755 255868

Rsym (%) 6.7 (22.2)* 23.0 (64.0) 20.7 (69.0) 11.4 (49.6) 11.0 (47.7) Completeness (%) 93.2 (76.4) 82.2 (79.3) 91.0 (85.2) 93.5 (94.1) 96.0 (92.3) Mean I/σ (I) 13.8 (2.6) 3.4 (1.0) 4.3 (1.0) 5.3 (1.6) 5.1 (1.4) Wilson B factor (Å2) 20.3 42.0 33.5 16.2 24.8 Refinement

R / Rfree (%) 16.8 / 21.4 23.4 / 31.9 23.4 / 29.1 20.7 /25.4 22.2 / 26.7 Monomers/asymmetric 4 4 4 4 4 unit No. of atoms Protein (chain A,B,C,D) 32660 32660 32660 32660 32660 (4100 res.) Ligand - 312 44 48 48 Waters 3766 836 1578 1427 923 Other atoms 130 (15 GOL, 42 (7 GOL) 70 (12 GOL, 4 Cl-) 48 (8 GOL) 30 (5 GOL) 9 OAc, 2 Cl-) B factors (Å2) main chain (A;B;C;D) 26.7; 36.4; 33.2; 39.1; 42.9; 37.4; 42.8; 48.1; 39.5; 17.0; 29.5; 29.1; 30.0; 41.0; 40.6; 28.8 44.8 53.8 18.7 31.6 ligand 38.1; 42.8; 41.6; 34.8; 36.5; 35.4; 28.0; 42.0; 38.9; 36.7; 44.7; 44.0; 39.3 36.1 41.7 47.6 ligand2 47.0; 66.8; 42.0; 72.2 Waters 39.1 29.5 46.0 32.0 30.2 Geometry RMSD Bond lengths (Å) 0.007 0.009 0.010 0.014 0.011 RMSD Bond angles (o) 1.1 1.3 1.3 1.5 1.3 Ramachandran favored 96.5 91.9 95.0 96.0 95.7 (%) Ramachandran outliers 0.2 0.5 0.2 0.2 0.1 (%) Molprobity score 1.95 2.95 2.34 2.05 2.21 PDB accession code 2x2h 2x2i 2x2j 4amw 4amx

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The obtained structure of native GLase was used for molecular replacement with PHASER to solve the structures of GLase in complex with 1-deoxynojirimycin (GLase- DNJ), acarbose (GLase-Acr), 5FαGlcF and 5FβIdoF. In the GLase-DNJ structure one deoxynojirimycin molecule was observed in the active site of each GLase molecule in the asymmetric unit. Continuous density in Fo - Fc maps for 2 acarbose molecules/protein molecule was observed in the GLase-Acr crystal, one bound in the active site and one bound at a remote location. In the secondary substrate binding sites electron density is only visible for three of the four sugar residues; the reducing end glucose points into the solvent and is probably flexible. After rigid body refinement using REFMAC5 2Fo – Fc and

Fo – Fc maps clearly showed density for 5FαGlc and 5FβIdo bound in the active sites. The inhibitor molecules were built with COOT and the structures were refined using REFMAC5. No significant conformational changes are observed between native and ligand-bound enzymes. The Cα atoms of GLase superimpose with root mean square deviations of 0.3, 0.2, 0.3 and 0.2 Å for GLase-Acr, GLase-DNJ, 5FαGlc and 5FβIdo, respectively. Analysis of the stereochemical quality of the models was done with MolProbity (Chen et al., 2010). Figures were prepared with PyMOL (Schrödinger).

Sulfolobus solfataricus α-Glucosidase (MalA) Mutagenesis, Expression, Purification and Characterization

Wild type and mutant strains of MalA, with codon optimization for growth in Escherichia coli, were ordered from Sloning BioTechnology (Puchheim, Germany) and were inserted into the isopropyl-(β)-D-thiogalactopyranoside (IPTG)-inducible expression vector pET-

24a which contains a kanamycin resistance marker and a C-terminal His6 tag. For the construction of the Glu323 mutant primers containing the mutation were introduced with the QuikChange Site-directed mutagenesis kit, using plasmid pET-24a-MalA- I213V/I249N/D251T (triple mutant) as the template. Successful mutagenesis was confirmed by nucleotide sequencing. Expression of the construct was achieved in E. coli BL21(DE3) cells grown in LB medium. All cultivation media contained 50 μg ml-1 kanamycin. At an A600 of 0.5-0.7, the cells were induced with 1.0 mM isopropyl-β-D- thiogalactopyranoside for a few hours at 180 rpm and 37 °C. SDS-PAGE was performed to investigate protein expression. The cell pellets were resuspended and washed in 50 mM Tris–HCl, pH 8.0 with 100 mM NaCl and disrupted with a French press. After centrifugation, the resultant extract was partly purified by HisTrap HP (GE Healthcare) chromatography with an imidazole gradient in 20 mM Tris, pH 7.5. Further purification was achieved by gel filtration chromatography using a Superdex 200 column with 20 mM Tris, pH 7.5, and 0.2 M NaCl. α-Glucosidase activity was analyzed as in (Ernst et al., 2006). Glucan lyase activity was assayed with the DNS reagent (Yu et al., 1998) and with O-ethylhydroxylamine at 21 °C (Hirano et al., 2000)

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After the reaction of enzyme with maltose, the samples were incubated with O- ethylhydroxylamine/HCl at 21 °C for at least 2 h and then centrifuged at 13,000 rpm. The resulting oximes were separated on an ÄKTAmicro system (GE Healthcare) equipped with a Sephasil C18 analytical column (4 ⨯ 250 mm, 5 μm;GE Healthcare) with as mobile phase an increasing gradient of acetonitrile (0–50%) in water,pumped at a flow rate of 0.3 ml/min. Detection was performed with a UV monitor at 207 nm. Pure anhydrofructose at different concentrations was used as a positive control.

RESULTS AND DISCUSSION

Overall Structure of α-1,4-Glucan Lyase

The structure of full-length Gracilariopsis lemaneiformis GLase missing 11 residues at its N-terminus was solved by molecular replacement and refined against 2.06 Å resolution diffraction data to an R-factor of 0.168 (Rfree = 0.214) with good stereochemistry (Table 1, Fig. 3). Although the P1 unit cell contains 4 molecules of 1025 residues each, in solution the protein is monomeric, as shown by native PAGE analysis (Yu et al., 1993).

Figure 3. Domain structure of GLase shown in two different orientations. The individual domains are colored as indicated: N-terminal domain (domain N), yellow; central catalytic (β/α)8 barrel (domain A), red; subdomain B

(loopAβ3→Aα3), magenta; subdomain B' (loopAβ4→Aα4), orange; proximal C-terminal (domain C), blue and the distal C-terminal domain (domain D), green. The catalytic residues are displayed as sticks in cyan and labeled as D553 and D665

Each GLase molecule contains four major structural domains, which form a very compact structure with dimensions of 93 ⨯ 65 ⨯ 49 Å (Fig. 3). GLase starts with the N-terminal domain (domain N, residues 12-332), and continues with the catalytic A domain

(residues 333-785). The A domain has a (β/α)8-barrel fold, with two inserted subdomains, B and B’. The structure is completed with the proximal and distal C-

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terminal domains (domain C, residues 784-879, and domain D, residues 880-1038, respectively). Domains N, C, and D are rich in β-sheets. The β-sheet content of GLase (28%) agrees with circular dichroism studies (Ernst et al., 2005). Despite the presence of 15 cysteine residues in GLase none of them forms disulfide bonds. The compactness of the GLase structure may explain its resistance to proteolytic cleavage. Only after prolonged reaction time, were proteinase K and subtilisin able to produce two fragments (Ernst et al., 2005), with the higher molecular mass fragment starting at Ser175, showing that the only site susceptible to these proteases is the surface loop 173ASSG176 (marked orange in supplemental Fig. S1). The GLase structure is very similar to the structures of the eight other structurally characterized GH31 family members, with DALI Z-scores over 30, despite root mean square deviation values >2.7 Å and sequence identities below 21%. These proteins are the N- and C-terminal catalytic subunits of human intestinal MGAM (NtMGAM; PDB code 2QLY (Sim et al., 2008), CtMGAM; PDB code 3TON (Ren et al., 2011)), human sucrase- isomaltase (SI; PDB code 3LPO (Sim et al., 2010)), Ruminococcus obeum α-glucosidase (Ro-αG1; PDB code 3N04 (Tan et al., 2010)), Sulfolobus solfataricus α-glucoside hydrolase (MalA; PDB code 2G3M (Ernst et al., 2006)), Cellvibrio japonicus α-xylosidase (CjXyl; PDB code 2XVG (Larsbrink et al., 2011)) and α-transglucosylase (CjAgd; PDB code 4B9Y (Larsbrink et al., 2012), and Escherichia coli α-glycosidase (YicI; PDB code 1WE5 (Lovering et al., 2005)). Most of the conserved residues are located in the A-domain (supplemental Fig. S1), in agreement with its important substrate-binding and catalytic function.

Domain N Contains a Secondary Carbohydrate Binding Site (SBS)

The large N-terminal domain folds into a double-layered twisted 19-strand β- supersandwich (Fig. 3). This fold belongs to the galactose-mutarotase-like superfamily and possibly functions as a carbohydrate binding domain in enzymes acting on saccharides (see the SCOP database (Murzin et al., 1995)). The N domain contains loops that interact intimately with the catalytic domain and accommodate several invariant residues crucial for activity and active site architecture (supplemental Fig. S1). Among 212 215 239 266 them are residues Gly , Glu , Asp , and Tyr from the NI, NII and NIII motifs GLAE, YNVD, and PMYYAAP, respectively (Ernst et al., 2006) (supplemental Fig. S1). Residue Gly212 has main chain torsion angles disallowed for other amino acids ( and  are 115° and 129°) nor is there space for a larger side chain. Residue Glu215 is hydrogen-bonded with one of its side chain oxygen atoms to the backbone amide of residue Tyr266, of which the hydroxyl group, in turn, makes a hydrogen bond with the backbone amide of residue Asp665, the catalytic acid. Residue Asp239 directly participates in substrate binding (see

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Crystal structure of α-1,4 glucan lyase ......

below). Thus, residues Gly212, Glu215 and Tyr266 appear important to stabilize the architecture of the active site in domain A. Expression of full-length G. lemaneiformis GLase in H. polymorpha yielded an active glucan lyase missing 11 residues at the N-terminus, as revealed by N-terminal amino acid sequencing. Nevertheless, its activity and substrate specificity were indiscernible from the wild type enzyme (Bojsen et al., 1999a, Yu et al., 1993), indicating that the 11 N- terminal residues were not essential for activity and expression in H. polymorpha. In contrast, no expression of variants with a larger N-terminal truncation (containing only residues 224-1038 or 224-938) was observed. Most likely no active enzyme is produced because these constructs lacked 2 of the 3 residues that stabilize the active site architecture. GLase binds to raw starch (Yu et al., 1993), but amino acid sequence alignments with enzymes having carbohydrate binding modules (CBMs) did not reveal any similarity (Yu et al., 1999a). The crystal structure of GLase-Acr revealed that GLase binds 2 acarbose molecules, 1 in the active site (see below) and the other to the N-terminal domain in a cleft between residues 32-37 and residues 189-191 (Fig. 4C). This latter acarbose is bound to the enzyme with its nonreducing end valienamine and dideoxyglucose units only, but not with its maltose part. Soaking experiments with the monosaccharide analog deoxynojirimycin (GLase-DNJ) and the fluorosugars 5FβIdoF and 5FαGlcF did not reveal any binding to the N domain of GLase, suggesting that minimally a α-1,4-linked glucose disaccharide is needed for recognition and binding by the N domain. Several glycoside hydrolase families are known to have CBMs associated with a starch binding function. Domain N of GLase differs from these known CBMs in its amino acid sequence, size, and fold, and its specificity for nonreducing end glucan disaccharide moieties. Aromatic amino acid are usually essential for the function of CBMs as they provide pi stacking interactions with the sugar rings (Cuyvers et al., 2012), but as seen in Fig. 4C, residues Phe33 and Trp41 in GLase bind acarbose via Van der Waals interactions. This implies that this binding site should be regarded as a SBS (Cuyvers et al., 2012). Sequence alignments of members of the GH31 family indicate that such SBSs exist in other algal glucan lyases, and a variant form may exist in fungal glucan lyases (Yu et al., 1999a). Although these findings suggest that domain N of GLase contains a SBS, it is not conserved in other GH31 members. A possible reason could be that such a binding site is only advantageous for proteins that act on larger substrates such as glycogen and starch granules.

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Figure 4. The active site. (A) GLase in complex with 1-deoxynojirimicyn (GLase-DNJ)(green) bound in subsite -1 with Fo-Fc omit electron density contoured at 3.0 σ. (B) GLase in complex with acarbose (GLase-Acr)(green) bound in subsites –1, +1, +2 and +3 with Fo-Fc omit electron density contoured at 3.0 σ. Further color coding of main chain and side chains are as in Fig 3. For clarity, only side-chains in panel B are shown. (C) Image of acarbose (green) bound in the putative starch-binding site at the N-terminal domain N of GLase-Acr in cpk coloured stick and transparent surface presentation.

Domains C and D

The proximal C-terminal domain consists of an eight-stranded antiparallel β-sandwich with a 310-helix and a small α-helix (αC1) inserted after the first strand. It is packed between the N, A and D domains (Fig. 3 and supplemental Fig. S1). The 6-residue 310-

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Crystal structure of α-1,4 glucan lyase ......

helix between βC5 and βC6, absent in other GH31 enzymes, is very well stabilized between the A and D domains. The 159 C-terminal residues of GLase form domain D. Domain D is made up of a 12- stranded mostly antiparallel β-sandwich, consisting of two sheets of six strands, two α- helices after strands βD3 and βD9, and two 310-helices after strands βD1 and βD11. This domain has major interactions with domains C and A, especially via helices αD1 and αD2 (supplemental Fig. S1). Expression of a truncated GLase variant (1-938) missing 100 residues from the C-terminus showed activity, although 10-20-fold decreased, indicating that the 100 C-terminal residues were not essential for activity and expression in H. polymorpha. Other GH31 members contain a similar D domain (see e.g. Nakai et al., 2006).

Domain A and the Active Site Structure

The catalytic domain A consists of a (β/α)8-barrel (TIM-barrel) with two additional helices, αA’ and Aα8’. The two additional domains, B (residues 462-530) and B’ (residues 556-615), are inserted after βA3 and βA4 respectively (Fig. 2 and supplemental Fig. S1). The secondary structure elements of subdomain B (a 3-stranded sheet and an α-helix) are the same as those in the other GH31 members NtMGAM, CtMGAM, Ro-αG1, SI, CjAgd and MalA, but in GLase the sheet is longer and closer to the active site. An extra loop in subdomain B (residues 490-504) interacts with the N-terminus and subdomain B’.

The active site of GLase is located at the C-terminal end of the (β/α)8-barrel of domain A. Residues from the loops following strands βA1, βA6, βA7 and βA8 shape the active site. Furthermore, loops from the N domain (243-262), the B domain (512-518), and B’- domain (559-590) make up the entrance to the active site (Fig. 2). The catalytic nucleophile Asp553 (Lee et al., 2002) is positioned at the end of strand βA4. The distance between the carboxylic acid oxygen atoms of residues Asp553 and Asp665 is 5.8 Å, which makes residue Asp665 the probable acid/base catalyst. This distance is similar to that found in retaining glycoside hydrolases, which is ~5.5 Å (McCarter et al., 1994). In the structure of native GLase, two glycerol molecules from the cryoprotectant, known to often mimic sugar substrates, are bound in the active site. The nucleophile Asp553 is hydrogen bonded to residues Arg649, Asn459, the O2 atom of a glycerol molecule and 2 water molecules. The acid/base Asp665 has interactions with 3 water molecules and the O1 atom of the second glycerol molecule. The residues at the C termini of the β-sheets in the active site vicinity include Phe373, Asp412, Asn459, Asp553, Arg649, Trp662, Asp694 and His731 from β-strands A1 to A8 respectively. Together with residues Trp551, Met554, Phe698, Arg729 and Tyr513, they form subsite –1. The narrow entrance to the –1 subsite, creating a true pocket, is mainly formed by residues Phe373, Tyr513, Met554, Arg649, Asp665 and Phe698. The overall contours of the active site pocket of

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GLase are similar to those of the GH31 members MalA, NtMGAM, CtMGAM sucrase- isomaltase, Ro-αG1, CjAgd, CjXyl and YicI. However, amino acid subsitutions of residues Phe373, Val413, Asn459, Thr461 and Tyr513 are observed (supplemental Fig. S1). Residue Tyr513 resides in subdomain B and is part of the extra loop, which is significantly different from a Trp residue that resides at the same site in other glycosidase GH31 members. Asn459 makes a hydrogen bond with the adjacent nucleophile Asp553. This hydrogen bond is absent in the glycosidase members of GH31 where an Ile residue or a smaller Ser residue is present.

Binding of 1-deoxynojirimycin and acarbose

1-Deoxynojirimycin, a glucose analogue with a nitrogen instead of an oxygen atom in the pyranose ring, is bound in subsite –1 via hydrogen bonds (Fig. 4A). The hydroxyl groups of the 1-deoxynojirimycin make hydrogen bonds to residues Asp665, Arg649, His731, Asp412, Asp553, Asn459 and 2 water molecules. Residues Phe698, Phe373, Tyr513, Trp551 and Trp662 provide hydrophobic interactions. Acarbose is bound to subsites -1 to +3 of GLase, with the valienamine residue binding to subsite -1, the 6-deoxyglucose residue to subsite +1 and the maltose subunit to subsites +2 and +3. Acarbose has a somewhat twisted conformation at the N-glycosidic bond destroying the hydrogen bond between the sugar OH2 group in subsite –1 and the sugar OH3 group in subsite +1 as shown in Fig. 4B. Interactions involving the valienamine moiety of acarbose at subsite –1 are similar to those exhibited in the complex of 1- deoxynojirimycin and GLase. The nitrogen atom bound at C1, which mimics the glucosidic oxygen atom in α-1,4 linkages, is located between the two catalytic aspartates. Residue Asp665-OD2 makes a hydrogen bond to the nitrogen atom, which would be an ideal orientation for protonation of a glucosidic oxygen. The binding of acarbose in subsite -1 is surprisingly similar to that of acarbose binding in NtMGAM, CtMGAM and CjAgd.

Carbohydrate Binding Subsites +1, +2 and +3

For subsite +1, residues Arg649, Asp239 and Met553 interact with the OH3 group of the 6- deoxy-glucose unit of acarbose. The OH2 group has hydrogen bonds with OD1 and OD2 of residue Asp239. These hydrogen bonds compensate for the loss of the hydrogen bond between the OH2 group of the valienamine moiety and OH3 of the 6-deoxyglucose unit of acarbose. Residue Asp239 is conserved among GH31 members and always resides in the N-terminal domain. These interactions are essentially similar to those seen in the NtMGAM-acarbose complex. Mutating the equivalent residue of Asp239 in Ro-αG1 eliminated enzymatic activity, indicating its importance (Tan et al., 2010). In CjXyl and

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Crystal structure of α-1,4 glucan lyase ......

CjAgd the role of Asp239 is taken over by a Glu, present in the PA14 domain insert in the N-terminal domain of CjXyl and in the B’ domain of CjAgd (supplemental Fig S1). At the position matching the absent OH6 group of the dideoxyglucose unit enough space is available for a hydrogen bond of O6 to the backbone carbonyl of residue Tyr513 when an α-1,4-glucan is bound. The +1 sugar ring stacks against a hydrophobic cushion formed by the side chain of residue Tyr513 (Trp in the other GH31 family members) and residue Met554 (Met in other members, except for Ala in CjXyl, Leu in CjAgd and Phe in YicI) (supplemental Fig. S1). A Trp residue at position 513 is strictly conserved in glycoside hydrolase members of family GH31 whereas in GLase it is replaced by a Tyr residue (Yu et al., 1999a). The side chain of residue Leu241 is involved in hydrophobic interactions with the sugar rings at subsites +1, +2 and +3 (Fig. 4B). At subsite +2, residue Gln245 interacts with OH2 group of the glucose unit of acarbose and residue His741 on the opposite site of the groove interacts with OH6 group. At subsite +3, there are no interactions between the substrate and GLase (Fig. 4B). The binding of acarbose completely shields subsite -1 from the solvent. Whereas residue Thr205 in NtMGAM and residue Gly89 in MalA allow space for entering water molecules used in the hydrolytic reaction, in GLase residue Leu241 blocks the entrance of water molecules (Fig. 4A). All hydrolases from family GH31 have a small or hydrophilic amino acid at this position, whereas the lyases have a larger hydrophobic amino acid (Leu or Met). Although sucrase-isomaltase also has a leucine at this position, it is located 2.5 Å away from Leu241, thus creating more open +1 to +3 subsites. The α-transglucosylase, CjAgd, has also strong interactions with the acarbose, via a hydrophobic “sugar” clamp formed by 2 tyrosine residues (Larsbrink et al., 2012). A similar clamp, made by 2 Trp residues, has also been observed in CjXyl (Larsbrink et al., 2011). The binding of 1- deoxynojirimycin and acarbose, inhibitors of GLase, is illustrated in the second image of Scheme 1.

Substrate Specificity of GLase

Compared with the GH31 α-xylosidases YicI and CjXyl subsite -1 of GLase has more space enabling it to accommodate the hydroxymethyl group of α-1,4-glucans. Indeed, the double mutation C307I/F308D of YicI, designed to increase the space in the active site, converted the α-xylosidase into an α-glucosidase (Okuyama et al., 2006). Furthermore, the carbohydrate-binding +2 subsite present in GLase allows the enzyme to act on longer α-1,4-glucans, in contrast to MalA, where subsite +2 is absent, and which consequently prefers the short disaccharide substrate maltose. Finally, the GH31 glucoside hydrolase Ro-αG1 prefers α-1,6- over α-1,4-linked substrates. This preference can be switched by a single W169Y mutation, which makes room for the 6-hydroxyl group of an α-1,4-linked glucose bound in subsite +1 (Tan et al., 2010). In GLase the equivalent residue is a

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phenylalanine (Phe373), which also provides space for the 6-hydroxyl group of the +1 subsite sugar residue, in agreement with the strict preference of the enzyme for α-1,4- linked glucosides.

5FαGlcF and 5FβIdoF Bind Covalently to GLase

Soaking GLase crystals with the mechanism-based inhibitors 5FαGlcF and its C5-epimer 5FβIdoF revealed that these compounds were covalently bound via a β-glycosidic bond to Asp553 in three of the four monomers. However, in monomer D they have reacted because no electron density was present for a covalent bond to Asp553 and for a fluorine atom at C1 (Fig. 5). Indeed, the product 5F-anhydrofructose (or its C5 epimer) with a planar O5-C1-C2-O2-C3 conformation could be fitted in the density present in the active site of monomer D. At the current resolution it is not possible to distinguish between the 1,2 enol and the keto forms of the product but we have modelled the keto-5F- anhydrofructose (Fig. 5).

Figure 5. Stereo view comparing the structures of 5FαGlc in the covalent intermediate (green/cyan) for monomer A, 5F-anhydrofructose/product (orange) in monomer D and 5F-anhydrofructose/product in monomer of 5FβIdo (slateblue).

5FαGlc and 5FβIdo differ in the fluorine position at C5. In the glucose sugar the fluorine atom is positioned axially and has Van der Waals interactions with Phe698. The glucose O6 hydroxyl is at hydrogen bonding distance to the OD2 atom of Asp412. In contrast, the fluorine atom in the idose sugar is equatorially positioned and interacts with ND2 of Asn459. The O6 of the idose sugar makes hydrogen bonds to a glycerol molecule and a water molecule. The different hydrogen-bonding interactions of the idose sugar may explain the higher affinity and inhibitory power of this compound toward the enzyme (Lee et al., 2002).

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Crystal structure of α-1,4 glucan lyase ......

Figure 6. Stereo view of simulated annealed omit electron density (Adams et al., 2010) of 5FβIdo (yellow) bound to Asp553. The density is shown for monomer A and is contoured at 3σ. For clarity, only the side-chains are shown that have interactions with the fluorine atom and C2-OH hydroxyl group of the covalently bound sugar.

The covalently bound fluorosugars have similar interactions as deoxynojirimycin except 4 for their C1 and C2 atoms. Their conformations are distorted from the normal relaxed C1 1 conformation (with an equatorial glycosidic bond) to a S3 skew boat conformation, in which the glycosidic linkage is pseudo-axial (Figs. 5 and 6). The skew boat conformation allows a closer approach of the OD1 carboxyl oxygen of Asp553 to the C2 atom of the -1 4 glucose which would not be possible with the C1 conformation of glucose (see third image of Scheme 1). This conformational distortion and the short distance between the carbonyl oxygen and the glucose C2 atom have important consequences for the catalytic mechanism (see below). The 5FαGlc inhibitor is bound in the same conformation as in CjAgd (PDB code 4BA0 (Larsbrink et al., 2012)). In both structures the trapped 5FαGlc molecules are covalently linked to the OD2 of the nucleophiles. One remarkable difference in the -1 pocket is Asn459, making a hydrogen bond to Asp553, whereas in CjAgd this residue is a hydrophobic isoleucine as in all other glycosidase members of GH31.

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Catalytic Mechanism

The crystal structures of GLase with various bound inhibitors indicate that the substrates bind with their nonreducing end in subsite –1, and extend toward the +n subsites. This binding mode is in agreement with biochemical experiments indicating that GLase degrades polysaccharides from the nonreducing end (Yu et al., 1999a). 1- Deoxynojirimycin and acarbose are known to inhibit glucan hydrolases by mimicking the oxocarbenium ion-like transition state structure (Lee et al., 2002). In glucan lyase they may function in the same way because the first step in the reaction, the formation of a covalent enzyme-substrate intermediate, which proceeds via an oxocarbenium ion-like transition state, is the same for GH31 lyases and hydrolases (3, 5). In GLase, this first step is catalyzed by the Asp665 side chain, which acts as a general acid catalyst protonating the glycosidic oxygen of the scissile bond. Concomitantly, one of the carboxylate oxygen atoms of the nucleophile Asp553 attacks the C1 carbon of the substrate’s glycosyl residue bound at subsite –1, generating a covalently bound glycosyl-enzyme intermediate (Scheme 1 and Figs. 5 and 6). For the second reaction step, the deglycosylation, Lee et al. (Lee et al., 2003) have suggested an E1-like E2 concerted mechanism on the basis of kinetic isotope effect measurements. This means that, at the anomeric centre, the glycosidic C-O bond is largely broken, generating a species with a highly developed oxocarbenium ion character, thereby acidifying the C2 proton and allowing its relatively facile removal late along the reaction coordinate. As suggested by the structures with covalently bound 5FαGlcF and 5FβIdoF the carbonyl oxygen atom of the nucleophile Asp553 is correctly positioned to abstract the proton from the sugar C2 carbon atom. Our proposed mechanism is similar to that of retaining glycosidases catalyzing the hydration of glycal substrates by syn-addition (the reverse of syn-elimination) of the C2 proton. In these enzymes the nucleophile is also assumed to participate in proton transfer (Lee et al., 2005, Trincone et al., 2001). Both the acid/base catalyst Asp665 and the nucleophile Asp553 have, respectively, been assumed to have a dual role in proton abstraction by Yu et al. (Yu et al., 1999a) and by Yip and Withers (Yip et al., 2006). Also, the possible existence of a separate base B- in the active site (Lee et al., 2005) or a water molecule activated by Asp665 (Yu, 2008) have been suggested. The acid catalyst Asp665 is positioned on the opposite side of the ring and consequently cannot abstract the proton from C2, nor is there a suitable water molecule situated. A separate base in the active site, able to abstract the proton, has not been observed. Arg649 makes a hydrogen bond to O2 of the sugar and is also not correctly positioned. Asn459 is properly located but with 4.6 Å too far from C2 to abstract the proton. Instead, Asn459 donates a hydrogen bond to the adjacent carboxylate of nucleophile Asp553 (ester oxygen). In monomer D of the 5FαGlcF experiment this bond

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Crystal structure of α-1,4 glucan lyase ......

becomes shorter once the covalent intermediate has been released. Asn459 probably plays a key role in the second transition state stabilization by modulating the position and charge of the nucleophile.

Scheme 1. Proposed reaction mechanism of α-1,4-glucan lyase, modified from Yip & Withers (Yip et al., 2006).

Arg649 is strictly conserved in all GH31 enzymes. In GH31 hydrolase members this arginine has a salt bridge interaction with a conserved Glu at position 556 (supplemental Fig. S1), but in the GH31 lyases the residue equivalent to Glu556 is a Val or Thr. In GLase Arg649 makes hydrogen bonds with the OH3 atom of the glucose bound in subsite +1 and the OD1 atom of the catalytic nucleophile Asp553. These interactions suggest that Arg649 is not only important for substrate recognition, but may also play a role in modulating the ionization state of Asp553. The arginine side chain is in an apolar environment conferred 266 662 554 556 by the side chains of Tyr , Trp , Met , and Val , which lowers the pKa of its guanidinium group (Guillen Schlippe et al., 2005), enabling it to pick up the proton abstracted from the sugar by Asp553. Subsequently, this proton can be relayed to the solvent via Asp239 or to the catalytic acid Asp665 for the next catalytic cycle after the product has left the active site. Indeed, in monomer D of the 5FβIdoF experiment the Arg649 guanidinium group has rotated by approximately 25o toward Asp553 OD1 thereby reducing the distance to Asp553 OD1 from 3.4 to 2.9Å. This suggests that residue 556

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(being either a Glu or a Val) is a major determinant of the hydrolase vs lyase reaction specificity of GH31 enzymes.

Shift of α-glucoside hydrolase activity to lyase activity

As argued above, the nature of the amino acid residue at position 556 (Val/Thr versus Glu) is very important for the reaction and product specificity of GH31 enzymes. Support for this notion comes from kinetic studies of the GH31 α-glucosidase from Schizosaccharomyces pombe (Okuyama et al., 2001), which showed that mutation of the residue equivalent to Val556 of GLase (E484A) eliminated its hydrolase activity, but that the enzyme retained 5% hydration activity on D-glucal to produce 2-deoxy-D-glucose by protonation at C2 (the reverse reaction of GLase). To demonstrate the importance of the residue at position 556 for lyase activity, an E323V mutation (Val556 in GLase) was introduced into the archaeal GH31 α-glucosidase MalA. We used a triple mutant of MalA (I213V/I249N/D251T; GLase Val413Asn459Thr461), which already had greatly reduced hydrolase activity, but was unable to form anhydrofructose. The specific activity of the partially purified triple mutant was 0.012 ± 0.004 μmol/min/mg of protein, compared with 0.362 ± 0.005 μmol/min/mg of the partially purified wild type MalA. For the quadruple MalA mutant (I213V/I249N/D251T/E323V) the expressed protein formed (soluble) aggregates after HisTrap purification. By storing the protein at 22 oC and removing the imidazole, a MalA protein of about 450 kDa could be resolved by gel filtration, which is similar to the molecular mass described previously (Ernst et al., 2006). The quadruple mutant showed detectable lyase activity at high protein concentration and after prolonged reaction time (Fig. 7), whereas it had completely lost glucosidase activity. Thus, the E323V mutation in MalA shifts the reaction toward lyase activity demonstrating the importance of position 323 (556) for hydrolase/lyase activity, likely by modulating the conformation of the fully conserved arginine in the active site.

Evolving MalA into a genuine lyase with a kcat similar to GLase will require further mutational studies.

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Crystal structure of α-1,4 glucan lyase ......

Figure 7. Chromatogram of the reversed-phased HPLC analysis. Indicated are the reaction products of GLase and MalA quadruple mutant on maltose, AF is the pure 1,5-anhydro-D-fructose. The sugars are derivatized with O- ethylhydroxylamine before subjecting to HPLC (Hirano et al., 2000). Reaction time for the quadruple MalA mutant was 90 hours, for wild type GLase 30 minutes at 21 oC. Calculations based on peak area of the HPLC analysis indicate that the MalA mutant has an activity of 0.029 nmol AF min-1 mg-1 and wtGLase 2.9 μmol AF min-1 mg-1. The quadruple mutant has 0.001% (~100,000 times lower) activity of the wtGLase at 21 oC. The retention time of the derivatized AF is 7.69 ml. Further detailed kinetic characterization of the MalA mutants remains to be done.

Structures of all four subgroups of GH31 (Ernst et al., 2006) have now been determined. Intriguingly, whereas the three-dimensional structures and catalytic site architectures are very similar and the essential catalytic residues are fully conserved, GLases show very different reaction specificity compared with the GH31 glucan hydrolases. Our results show that indeed the first step of the reaction, the formation of a covalent enzyme-substrate intermediate, is conserved. However, the glucosyl intermediate is not hydrolyzed like in the GH31 hydrolases, but a subtle change, a Val instead of a Glu at position 556, in the active site now promotes a lyase reaction. As a consequence the nucleophile in the lyase has acquired a dual function and can also act as a base, abstracting the proton from the C2 atom of the -1 glucose residue. Nature therefore has chosen a simple way in evolving new enzyme activities through clever changes at a localized area in the active site environment.

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ACKNOWLEDGEMENTS

We thank Ms K. Larsen and C. P. Walter for their excellent technical assistance with expression and characterization of the glucan lyase, and Dr H. Pedersen for help with the fermentation of the Hansenula strain. We are grateful to staff scientists and local contacts at beam lines ID14-4 and ID23-2 at the ESRF, Grenoble, France, and BW7A at the EMBL- Hamburg outstation (DESY), Germany, for their support during data collection.

SUPPORTING MATERIAL

Figure S1. Structure–based alignment of the red algal GLase, human N-terminal and C-terminal maltase- glucoamylase (NtMGAM and CtMGAM), human sucrase-isomaltase (SI), bacterial α-glucosidase (Ro-αG1), prokaryotic α-glucosidase (MalA), two bacterial α-xylosidases (YicI and CjXyl) and bacterial α-transglucosylase (CjAgd). The structural alignment was made with SSM (http://www.ebi.ac.uk/msd-srv/ssm/). For clarity reasons the trefoil Type-P domains of NtMGAM, CtMGAM and SI have been deleted. Also have been deleted the PA14 domain (res. 233-383) and loop701-717 of CjXyl. The secondary structure elements on top of the sequence alignment are placed with respect to the crystal structure of GLase. Identical residues have a red background color, homologous residues have a red color. Residues in the –1 subsite have a purple background, catalytic residues have a cyan background. Residues involved in binding acarbose in the N-terminal domain are colored green, the conserved regions in the N-terminal domain, NI, NII and NIII motifs, are labeled with blue triangles and the residue important for product specificity is labeled with a cyan triangle. The figure was created with ESPript (http://espript.ibcp.fr/ESPript/ESPript/) (Gouet et al., 1999).

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Chapter 4

Crystal structure of quinone-dependent alcohol dehydrogenase from Pseudogluconobacter saccharoketogenes a versatile dehydrogenase oxidizing alcohols and carbohydrates

Henriëtte J. Rozeboom*, Shukun Yu†, Rene Mikkelsen†; Igor Nikolaev†; Harm Mulder† and Bauke W. Dijkstra*

*Laboratory of Biophysical Chemistry, Groningen Biomolecular Sciences and Biotechnology Institute, University of Groningen, Nijenborgh 7, 9747 AG Groningen, The Netherlands. †DuPont Industrial Biosciences, Edwin Rahrs Vej 38, DK 8220 Brabrand, Aarhus, Denmark and †Archimedesweg 30, 2333 CN Leiden, The Netherlands

Manuscript in preparation

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ABSTRACT

The quinone-dependent alcohol dehydrogenase (PQQ-ADH, E.C. 1.1.5.2) from the Gram- negative bacterium Pseudogluconobacter saccharoketogenes IFO 14464 oxidizes primary alcohols (e.g. ethanol, butanol), secondary alcohols (monosaccharides), as well as polysaccharides and cyclodextrins. The recombinant protein, expressed in Pichia pastoris, was crystallized and 3D structures of the native form, with PQQ and a Ca2+ ion, and of the enzyme in complex with a Zn2+ ion and a bound substrate mimic were determined at 1.72 Å and 1.84 Å resolution, respectively. PQQ-ADH displays an eight- bladed β-propeller fold, characteristic of Type I quinone-dependent methanol dehydrogenases. However, three of the four ligands of the Ca2+ ion differ from those of related dehydrogenases and they come from different parts of the polypeptide chain. Remarkably, no vicinal disulfide bridge is present near the PQQ, which in other PQQ- dependent alcohol dehydrogenases is implicated in electron transfer. These differences result in a more open, easily accessible active site, which explains why PQQ-ADH can oxidize a broad range of substrates. The bound substrate mimic suggests Asp333 as the catalytic base.

INTRODUCTION

Quinoproteins are oxidoreductases containing an amino acid-derived o-quinone cofactor, such as pyrroloquinoline quinone (PQQ; also named Methoxatin), topaquinone (TPQ), Tryptophan tryptophylquinone (TTQ), lysine tyrosylquinone (LTQ), and cysteine tryptophylquinone (CTQ). The best-studied cofactor is PQQ, which is a heterocyclic compound with two quinone oxygen atoms, two heterocyclic nitrogen atoms, and three carboxylate oxygen centers. It binds non-covalently to proteins and can serve to bind metal ions. In enzymes so far crystallographically characterized, generally a Ca2+ ion is observed, coordinated by the PQQ O5, N6, and O7A atoms (Ghosh et al., 1995, Oubrie et al., 1999). Quinoproteins have been recognized as the third class of redox enzymes after nicotinamide- and flavin-dependent dehydrogenases (Hauge, 1964). They oxidize a wide range of saccharide, alcohol, amine, and aldehyde substrates, and the PQQ-containing quinoproteins have been classified into two classes on the basis of their primary and tertiary structures (Yamada et al., 2003). One class features a six-bladed ‘propeller fold’ and includes soluble glucose (sGDH) (Oubrie et al., 1999) and some aldose dehydrogenases (Southall et al., 2006, Sakuraba et al., 2010). The other class is characterized by an eight-bladed propeller fold, and comprises three distinct types of quinone-dependent alcohol dehydrogenases (Anthony, 2001, Toyama et al., 2004). Type

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I enzymes include methanol dehydrogenases (MDHs) and quinone-dependent alcohol dehydrogenases (QADHs). MDHs are tetramers with a α2β2 structure; the α-subunits contain the PQQ cofactor and a Ca2+ ion, while the β-subunit is very small and has no known function (Ghosh et al., 1995). They are mostly found in the periplasmic space of methylothrophic bacteria, where they convert methanol into formaldehyde. QADHs are mostly homodimeric enzymes found in Proteobacteria like Pseudomonas, Ralstonia and Comamonas species. They convert primary and secondary alcohols and aldehydes into the corresponding aldehydes/ketones or acids (Keitel et al., 2000). Type II enzymes are monomeric quinohemoprotein alcohol dehydrogenases (QH-ADHs), which convert the same substrates as Type I QADHs, but not methanol. They have two distinct functional domains of which one is a cytochrome c domain with a covalently bound heme c group (Oubrie et al., 2002, Chen et al., 2002, Toyama et al., 2005). Finally, Type III enzymes are multicomponent membrane-bound alcohol dehydrogenases present in acetic acid bacteria (Toyama et al., 2004). Membrane-associated glucose dehydrogenases (mGDH) (Yamada et al., 2003, James et al., 2003) and polyol dehydrogenases (Soemphol et al., 2008, Holscher et al., 2009) are also denoted as type III enzymes. The PQQ-dependent alcohol dehydrogenase (PQQ-ADH) from the Gram-negative Pseudogluconobacter saccharoketogenes IFO 14464 (E.C. 1.1.5.2) is a monomeric enzyme that was initially found to be able to oxidize L-sorbose to 2-keto-L-gulonic acid via L- sorbosone (Shibata et al., 2001). However, the enzyme also oxidizes other monosaccharides, primary and secondary alcohols, aldehydes, α- and β-cyclodextrin, oligosaccharides, and polysaccharides, including pectin, carrageenan, guar gum, alginate, and carboxymethyl cellulose, although the larger molecules are oxidized with less efficiency (Shibata et al., 2001, Mikkelsen et al., 2011). Maltotetraose (G4) and maltoheptaose (G7) can be oxidized at the reducing end C1, yielding the carboxylic acid, as well as at their C6 positions, which yields aldehyde as the major and carboxylic acid as the minor product (Mikkelsen et al., 2011). The enzyme’s broad substrate specificity makes it of interest for biocatalytic applications. In the present study we determined the X-ray crystal structure of PQQ-ADH from P. saccharoketogenes IFO 14464, in its native form and in a complex with a polyethylene glycol fragment. These structures reveal an altered, more open active site conformation compared to other quinone-dependent methanol/alcohol dehydrogenases, which explains its unusual broad substrate specificity. Moreover, the present work also resolves the ambiguity of the active site catalytic base in Type I PQQ-ADH.

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MATERIALS AND METHODS

Cloning, expression and purification

The gene encoding the Pseudogluconobacter saccharoketogenes PQQ-ADH, including its own signal sequence, was synthesized as a codon optimized fragment by GeneArt AG (Regensburg, Germany) and cloned into the pDONR/Zeo cassette via the Gateway® BP recombination reaction (Life Technologies) resulting in the entry vector pENTRY-ADH. To enable expression of PQQ-ADH in Pichia pastoris, the gene was cloned from pENTRY- ADH into pPIC2-DEST via the Gateway® LR recombination reaction. pPIC2-DEST was derived from the commercially available vector pPIC-DEST by removal of the signal sequence from this vector. The resulting plasmid, pPIC2-ADH was linearized by SalI digestion, enabling integration of the construct into the HIS4 locus of P. pastoris GS115 upon transformation. This vector contains the P. pastoris AOX1 promoter, allowing strong methanol-inducible gene expression. For production of PQQ-ADH, P. pastoris transformed with pPIC2-ADH was grown in a 2 liter B. Braun Biostat B fermentor, and expression of the enzyme was induced with methanol according to standard P. pastoris fermentation protocols (Life Technologies). After fermentation the major fraction of the expressed PQQ-ADH was found in the culture supernatant, with levels reaching 100-400 mg/l in 172 hours after the start of the methanol induction. The N-terminus of the mature protein AEPSKAGQSA was found to start at position 37 of the coding sequence as determined by Edman degradation and analysis on a Procise® cLC capillary 491 protein sequencing system (Applied Biosystems, Foster City, CA, USA). The culture supernatant was extensively dialyzed against 20 mM Tris-HCl, pH 7.5 (Buffer A). The recombinant PQQ-ADH was purified by anion exchange chromatography using a 60 ml Q-Sepharose HP (XK16/20) column (GE Healthcare, Uppsala, Sweden) using a gradient of Buffer A, and Buffer A containing 1.0 M NaCl as Buffer B. Active fractions eluting at 0.35 – 0.40 M NaCl were pooled. Reconstitution with PQQ was carried out at room temperature in 5 mM CaCl2, 20 mM Tris-HCl, pH 7.5. The pooled fractions were concentrated and washed with GF buffer (20 mM Tris, pH 8.5, 5 mM CaCl2 and 150 mM NaCl) by ultrafiltration to 6.7 mg protein ml-1. Further purification was achieved on a HiLoad Superdex75 PG XK26/60 (320 ml) (GE Healthcare) column. The main fraction, containing active PQQ-ADH with a molecular weight of 68 kDa, was concentrated to 9.5 mg protein ml-1. PQQ-ADH activity was determined as described (Shibata et al., 2001, Groen et al., 1986). The enzyme assays were carried out in a reaction mixture (1.0 ml) containing 1% substrate, 160 μM 2,6-dichlorophenolindophenol (DCIP), 50 μM phenazine methosulfate (PMS), 20 mM Tris-HCl, pH 7.5, and reconstituted enzyme solution, and incubated at

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Crystal structure of PQQ-ADH ...... room temperature. Oxidation of the substrate was monitored at 600 nm for 10 minutes in a UV/Vis spectrophotometer (Ultrospec 4300 pro; GE Healthcare).

Crystallization and X-ray data collection

Vapor-diffusion crystallization experiments were conducted using a Mosquito crystallization robot (Molecular Dimensions Ltd, Newmarket, England). In a typical experiment, 0.1 μl screening solution was added to 0.1 μl protein solution in a 96-well MRC2 plate (Hampton Research, Aliso Viejo, USA), with the reservoir wells containing 60 μl screening solution. The screening solutions used for the experiments were Structure Screen, PACT (both from Molecular Dimensions Ltd, Newmarket, United Kingdom), Wizard (Emerald Biosystems, Bainbridge Island, USA), and JCSG+ (Qiagen Systems, Maryland, USA). Crystals grew from solutions containing polyethylene glycol (PEG) and zinc acetate. Crystallization conditions were optimized using hanging-drop set-ups with 18 to 21%

(v/v) PEG550 MME and 20 mM Zn(Ac)2 in 100 mM PCB buffer (sodium propionate, sodium cacodylate and BIS-TRIS propane, pH 6.0 – 7.0) (Newman, 2004) as precipitant, and drops containing 2 μl protein solution (9.5 mg ml-1 in 20 mM Tris-HCl, pH 7.5, 3 mM

CaCl2 and 150 mM NaCl) and 2 μl reservoir solution at 295 K. Crystals were only obtained from conditions with Zn2+ concentrations above 20 mM. Monoclinic crystals grew in space group C2 with cell dimensions a = 128.0 Å, b = 87.2 Å, c o = 57.1 Å and β = 90.2 . With 1 monomer of 65 kDa in the asymmetric unit, the VM is 2.5 Å3/Da (Matthews, 1968) with a solvent content of 51%. Before data collection, crystals were soaked for 15 seconds in a cryoprotectant solution, consisting of 25% (v/v)

PEG550 MME and 20 mM Zn(Ac)2 in 100 mM PCB buffer, pH 7.0, directly followed by flash cooling. Data were collected in house at 110 K with a DIP2030H image plate detector (Bruker-AXS, Delft, The Netherlands) using Cu-Kα radiation from a Bruker-AXS FR591 rotating-anode generator equipped with Osmic mirrors. Data were 95% complete up to 1.81 Å resolution; completeness of higher resolution shells became lower because of detector geometry. All intensity data were processed using iMOSFLM (Battye et al., 2011) and scaled using SCALA from the CCP4 package (Winn et al., 2011). The PQQ-ADH crystal was hemihedrally twinned (the monoclinic β angle was close to 90 degrees (90.16o)) with a twin fraction α of 0.2 (meaning partial twinning). The twin operator was h, -k, -l (180o rotation around α). A summary of data collection statistics is given in Table 1.

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Table 1 Data collection and refinement statistics.

native G4 soak Wavelength (Å) 1.54 0.93 Space group C2 C2 Cell dimensions (Å, o) (a, b, c, β) 127.9, 87.2, 57.1, 90.16 128.2, 87.2, 57.0, 90.29 Crystal mosaicity (o) 0.73 0.90 Number of chains per A.U. 1 1

VM (Å3 Da-1) 2.45 2.45 Solvent content 0.51 0.51 Resolution range (Å) 30.5 – 1.72 (1.81-1.72) * 36.0 – 1.84 (1.93-1.84)

Rmerge (%) 8.5 (22.9)* 8.1 (27.1)

Rpim (%) 5.4 (20.9) 4.1 (12.3) Completeness (%) 88.9 (31.3) 100.0 (99.9) 〈I/σ(I)〉 11.4 (4.3) 14.6 (6.2) Reflections total (unique) 194837(59031) 269946 (54822) Multiplicity 3.3 4.9 Wilson B (Å2) 9.5 10.5 Refinement Total protein atoms 4278 4278 Amino acid sequence numbers 45 – 606 45 - 606 Mean B-factor protein 6.8 7.4 Number of water molecules 626 571 Mean B-factors solvent 16.9 18.4 Number of ions 1 Ca2+, 4 Zn2+, 1 Cl-, 1 PPI 7 Zn2+, 1 Cl-, 1PPI Mean B-factors ions 14.3, 13.7, 13.0, 15.6 15.1, 12.2, 10.7 Ligands 1 PQQ 1 PQQ, 1 PEG Mean B-factors ligands 16.9 12.0, 29.4

Rcryst/ Rfree (%) 12.8 / 14.9 16.1 / 18.5 R.m.s. deviation bonds /angles 0.010 / 1.4 0.006 / 1.1 Ramachandran plot (%) 96.3 / 3.2 / 0.5 97.0 / 2.5 / 0.5 (favored/additionally allowed/ disallowed

MolProbity Clash-score / percentile 5.5 / 93rd 1.3 / 100th

MolProbity score / percentile 1.55 / 89th 1.03 / 100th * Values in parentheses are for the highest resolution shell.

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Structure determination and refinement

The structure of PQQ-ADH was solved by molecular replacement with PHASER (McCoy et al., 2007) using a search model (Schwarzenbacher et al., 2004), composed of quinohemoprotein alcohol dehydrogenase from Comamonas testosteroni (26% sequence identity, PDB code 1KB0) (Oubrie et al., 2002) and quinohemoprotein alcohol dehydrogenase from Pseudomonas putida HK5 ADH-IIG (25% sequence identity, PDB code 1YIQ) (Toyama et al., 2005), generated by the FFAS server (Jaroszewski, L., Rychlewski, L., Li, Z., Li, W. & Godzik,A., 2005). The phases obtained from the model were improved with the ARP/wARP procedure (Langer et al., 2008) combined with manual model improvement with Coot (Emsley et al., 2010). Refinement was done with REFMAC5 (Winn et al., 2011) with automatic twin refinement and using TLS rigid body refinement (Painter et al., 2006). Further details of the refinement are listed in Table 1. For maltotetraose binding studies, a 9-month old crystal was soaked in a solution containing 100 mM maltotetraose (G4) for several minutes and immediately flash cooled in liquid nitrogen. Data were collected from a non-twinned crystal on beam line X11 (EMBL outstation at DESY, Hamburg, Germany). After rigid body refinement with REFMAC5 using the native structure as starting model, 2Fo – Fc and Fo – Fc maps showed some extra, metal ion-like density in the active site. To identify the ions, we calculated an anomalous difference Fourier map, with data collected at a wavelength of 0.93 Å (2.3 anomalous electrons for zinc, and 0.5 for calcium), using phases obtained from the model without any metal ions. Analysis of the stereochemical quality of the models was done with MolProbity (Chen et al., 2010). Figures were prepared with PyMOL (http://www/pymol.org). Atomic coordinates and experimental structure factor amplitudes have been deposited in the Protein Data Bank (PDB) with PDB IDs 4CVB and 4CVC for native and substrate-bound PQQ-ADH, respectively.

RESULTS AND DISCUSSION

Overall Structure

The crystal structure of the quinone-dependent alcohol dehydrogenase from Pseudogluconobacter saccharoketogenes IFO 14464 (PQQ-ADH) was determined at a resolution of 1.72 Å to final R/Rfree values of 12.8/14.9%. The final model comprises one protein chain of 562 residues, one calcium ion, one PQQ cofactor, three zinc ions, one chloride ion, one propanoic acid molecule and 614 water molecules. The first 8 residues of the mature protein (residues 37-44) and the last 2 residues were not visible in the electron density maps. Further details of the refinement are listed in Table 1. Most of the

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φ/ψ angles fall in the allowed regions of the Ramachandran plot; 96.3% are in the most favored regions, 3.2% in the additionally allowed regions, and 0.5% in the disallowed regions (Val196, Pro432 and Val475). Pro432 is forced into an unusual geometry by the disulfide bond between the adjacent Cys431 and Cys460. Val196, in a β-turn, and Val475, next to the active site calcium ion ligand Tyr476, are also forced by steric hindrance into the disallowed region. The three-dimensional structure of PQQ-ADH shows an eight-bladed β-propeller fold (Fig. 1). Each of the eight propeller blades (W1 – W8) contains 4 antiparallel β-strands (A-D) forming a twisted β-sheet (also called W-motif). After W5, three additional strands are inserted (2a, 2b and 3a), two forming a β-sheet, and one is part of a three-stranded sheet formed by the insertion of two strands in W6 after strand B (Figs. 1b and S1). In addition, a small α-helix (residues 579-583) is inserted after strand B of W8. All these features are also observed in QADHs and MDHs. Furthermore, an extra strand (W5E in Fig. S1, Fig. 1B) is inserted between W5A and W5B. This strand is a unique feature of PQQ-ADH. It contains a zinc-binding site important for crystal contacts (Fig. 2C) (see below).

Figure 1. Three-dimensional structure of Pseudogluconobacter saccharoketogenes PQQ-ADH. (A) Ribbon diagram showing the protein as a cartoon in rainbow colors from the N-terminus in blue to the C- terminus in red. (B) Ribbon diagram showing the protein rotated by 90° compared to (A) with helices colored red and the β-propeller in blue. The three-stranded sheet is colored purple and the two-stranded sheet cyan. Strand W5E (in teal) is a unique feature of PQQ-ADH. The PQQ cofactor is shown in grey sticks, the calcium ion as a green sphere, the chloride ion as a yellow sphere, the zinc ions as brown spheres and the disulfide bridges as sticks.

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Figure 2. Zinc-binding sites in PQQ-ADH (A-C) Amino acid residues and ligands of native PQQ-ADH are in shown in green, of symmetry related molecules in dark green and denoted with an asterisk (*). Zinc ions are shown as brown spheres, the chloride ion as a yellow sphere, and water molecules as red spheres. (D-E) Amino acids and ligands of the G4 structure in cyan. Zinc ions 1- 3 are of the native and zinc ions 5-7 are of the G4 structure.

Six of the eight β-propeller blades contain a conserved tryptophan-docking motif (Table 2). The invariant residues cause a tryptophan residue to stack between an alanine residue and the peptide bond of a glycine residue located on a neighboring β-sheet. The tryptophan-docking motif probably stabilizes the β-propeller fold (Keitel et al., 2000, Anthony et al., 1998).

Table 2. Tryptophan-docking motif Amino-acid sequences of the tryptophan docking motif of PQQ-ADH. The bold residues represent invariant or semi- invariant residues between motifs. The table has been adapted from (Shibata et al., 2001).

position 1 2 3 4 5 6 7 8 9 10 11 motif

W1 Ala127 Ile Asp Gly Lys Thr Gly Ser Leu Ile Trp137

W2 Ala178 Leu Asp Ala Lys Thr Gly Lys Leu Ala Trp188

W3 Gly230 Thr Asp Ala Glu Ser Gly Glu Glu Leu Trp240

W4 Ala312 Val Asp Pro Lys Thr Gly Glu Val Val Trp322

W5 Gln384 Phe Asp Ala Lys Thr Gly Asp Tyr Phe Trp394

W6 Ala496 Ile Asp Leu Ala Thr Gly Glu Thr Lys Trp506

W7 Ala537 Leu Asp Ala Glu Ser Gly Lys Glu Val Trp547

W8 Val601 Phe Ala Leu604 - Asp91 Leu Gln Leu Val Trp96 consensus Ala - Asp Ala Lys Thr Gly Glu Leu Val Trp

The 3D structure of PQQ-ADH is very similar to the propeller blade part of Type I MDHs (Xia et al., 1999, Williams et al., 2005, Nojiri et al., 2006, Li et al., 2011, Pol et al., 2014) and QADHs (Keitel et al., 2000), and Type II QH-ADHs (Oubrie et al., 2002, Chen et al., 2002, Toyama et al., 2005), with rmsd values of less than 1.7 Å and sequence identities around 27-30%. PQQ-ADH has the highest sequence and structural similarity to ADH-IIG from P. putida HK5 (Toyama et al., 2005) although this latter enzyme is a Type II quinohemoprotein and contains a cytochrome c domain. Unlike Type I MDHs and

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QADHs, PQQ-ADH is monomeric in solution. A phylogenetic reconstruction of the ten PQQ-containing alcohol dehydrogenases whose structures have been determined (see also Fig S1) shows that PQQ-ADH belongs to an outgroup and is more distantly related than the other enzymes.

Active site of PQQ-ADH

The active site is located near the upper face of the β-propeller in a hydrophobic cradle on the surface of the enzyme. Its solvent-exposed position enables it to accommodate a large spectrum of variously sized substrates. PQQ is bound via hydrogen-bonding interactions to the side chains of Glu106, Arg158, Ser203, Thr218, Gln220, Asp333, Glu335, Lys378, Asp439, Trp440 and Y515, and the carbonyl oxygen atom of Phe434 (Fig. 3). Additional binding interactions are provided via hydrophobic stacking interactions with Leu153 on one side of its molecular plane (see next paragraph) and with the indole ring of the invariant Trp270 on the other side. The reactive orthoquinone group is hydrogen-bonded to Asp439 and Lys378 with its O4 atom, and to the Lys378 side chain with its O5 atom. The O5 atom is also a ligand of the active site Ca2+ ion. The interactions of O5 have been suggested to polarize the C5–O5 bond (Zheng et al., 1997), priming the PQQ C5 atom to receive the hydride ion from the substrate, which is essential for the enzyme’s catalytic activity (see below).

Figure 3. Stereo image of the active site of PQQ-ADH. Amino acid residues in the active site are shown as green sticks, the PQQ cofactor as grey sticks and the calcium ion as a green sphere. Hydrogen bonds are indicated with black dashes and metal-ligand bonds with cyan dashes.

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As in other PQQ-dependent enzymes, the calcium ion is bound to the O5, N6 and O7A atoms of the PQQ cofactor. Additional coordination is provided by the side chains of Asp333, Gln220, Glu335 and Tyr476. The latter three ligands are unique to PQQ-ADH. To create a more open active site several active site loops have different conformations, and the calcium is coordinated by different residues. For instance, Glu335 replaces Asn255 as a Ca2+ ligand, and Gln220 replaces a Glu at a position equivalent to 221 in other quinoproteins. The Ca2+-coordinating Tyr476 side chain has no equivalent in the other PQQ-dependent alcohol dehydrogenases (Fig. S1). The solvent exposed and accessible active site suggests that a cytochrome c, the usual electron acceptor in the enzymes, could easily enter the active site to receive the electrons produced during the reaction. Superimposing the structure of PQQ-ADH on QH-ADH (Oubrie et al., 2002), which has a covalently attached cytochrome c domain, shows indeed that there is enough space in PQQ-ADH for a cytochrome c to bind (Fig. 4). Although PQQ-ADH has its broad substrate specificity in common with the PQQ- containing soluble aldose dehydrogenase from E. coli (Asd) (Southall et al., 2006), their sequence identity is only 7% and Asd belongs to the six-bladed β-propeller class of glucose and aldose dehydrogenases. The active site environment is also quite different between the two enzymes.

Figure 4. Structural comparison of the active sites of PQQ-ADH and C. testosteroni QH-ADH. Active site amino acid residues of PQQ-ADH are shown as green sticks. The PQQ cofactor is shown in grey sticks and the calcium ion as a green sphere. The amino acids residues, PQQ and calcium ion of QH-ADH are shown in magenta.

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Disulfide bridges

PQQ-ADH contains three disulfide bridges, Cys219-Cys226, Cys336-Cys377 and Cys431- Cys460. The Cys219-Cys226 and Cys336-Cys377 disulfides stabilize the positions of Gln220 and Glu335, respectively, which are both Ca2+ ligands. The Cys431-Cys460 disulfide is located between β-strands 3a and 3b and is also present in MDHs but not in QADHs. Surprisingly, PQQ-ADH does not contain a vicinal SS-bond between two adjacent cysteine residues in a Type VIII β-turn (Blake et al., 1994, Carugo et al., 2003) (Figs. 4 and S1). Such a vicinal disulfide is strictly conserved among quinohemo/quino alcohol dehydrogenases with the exception of lupanine hydroxylase, a type II QH-ADH (Hopper et al., 2003) and mGDH (Holscher et al., 2009). It has been shown that the disulfide bond is essential in electron transfer from PQQ to the recipient cytochrome c (Mennenga et al., 2009). An alternative suggestion is that the disulfide bond provides conformational rigidity of the protein during substrate binding (Oubrie et al., 2002). Instead of the vicinal disulfide bond, Leu153 interacts with the PQQ in PQQ-ADH (Figs. 3, 4 and S1), and the amino acid residues surrounding Leu153 adopt different conformations, creating a highly open and exposed active site. As evidenced by the crystal structure of PQQ-ADH, a vicinal disulfide bond is apparently not strictly required for enzyme activity of all PQQ- containing alcohol dehydrogenases.

Substrate Specificity of PQQ-ADH

PQQ-ADH (residues 37-606) shows broad substrate specificity, and oxidizes various primary and secondary alcohols, as well as various monosaccharides with good activity (Table 3). In solution, these monosaccharides exist predominantly in the pyranose form (Dworkin et al., 2000), and hence they are likely oxidized at their reducing end to the δ- lactone, which hydrolyzes in water to form the linear 1-acid end product. However, the monosaccharides are, to a small degree, also present in the open-chain aldehyde form (Dworkin et al., 2000). Therefore, we cannot fully exclude a direct oxidation of the aldehyde by PQQ-ADH to the 1-acid, in particular because the enzyme can oxidize linear monosaccharide alcohols like D-sorbitol (D-glucitol), xylitol and D-mannitol, albeit with somewhat lower activity (5-20%; Table 3). The enzyme’s activity on L-sorbose is less than 1% of the activity on glucose, although it is still well measurable. This low activity may be related to L-sorbose being a 2-keto-sugar. As is evident from Table 3, the enzyme also has significant activity on tri- and tetrasaccharides (20-30% of the activity on glucose). The activity on rac-2-methyl-2,4- pentanediol demonstrates the enzyme’s activity on secondary alcohols. The broad

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Crystal structure of PQQ-ADH ...... substrate specificity can be explained by the open active site, which is accessible for substrates of widely different sizes.

Table 3. Substrate specificity of PQQ-ADH

Substrate Relative activity (%) Sugars and sugar alcohols D-Glucose 100 D-Galactose 85 D-Mannose 77 D-Xylose 130 Methyl α-D-glucopyranoside 5 Xylitol 20 L-Sorbose 0.6 D-Sorbitol 9 D-Mannitol 6 Alcohols Methanol 3 Ethanol 16 Ethyleneglycol 109 n-Propanol 120 iso-Propanol 121 rac-1,2-Propanediol 128 Propan-1,2,3-triol (glycerol) 25 n-Butanol 241 rac-2-Methyl-2,4-pentanediol (MPD) 164 Oligosaccharides Maltotriose (G3) 31 Maltotetraose (G4) 24 * D-Glucose + 20 mM Zn(Ac)2 7 & G4 + 20 mM Zn(Ac)2 0.02 The enzyme assays were performed as described in Materials and Methods. The enzyme activity toward D-glucose * & was assigned as 100 (%). 96 h incubation time (with Zn(Ac)2); 3 h incubation time.

Zinc ions

Four zinc ions, originating from the crystallization solution, were found at the surface of PQQ-ADH (Fig. 2A-D). Two of them (Zn1 and Zn3) stabilize the interaction between symmetry related molecules in the crystal, rationalizing why PQQ-ADH crystals could not

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be grown in the absence of zinc ions. The other two zinc ions have no interactions with symmetry-related protein molecules. Three additional zinc ions were found in a 9-month old crystal used to investigate the binding of maltotetraose (G4) (Fig. 2E). An anomalous difference Fourier map showed, in total, seven strong peaks well above the noise level. The three extra Zn2+ ions probably result from the high concentration of ions and the prolonged storage time of the crystal. Unexpectedly, one of the strongest peaks was found at the position of the Ca2+ ion in the active site (Fig. 5). Refining this ion as a Zn2+ ion instead of Ca2+ resulted in a tetrahedral coordination arrangement (as preferred by Zn2+), with two metal-ligands (the Tyr476 and Asp333 side chains) moved out of the coordination sphere (Fig. S2). The remaining ligands refined to typical Zn2+-binding distances of 2.0-2.3 Å (Harding, 2001). Together, these data suggest that the calcium ion has been replaced by a zinc ion in PQQ-ADH-G4. Activity experiments with addition of 20 mM Zn2+ to the reaction medium showed decreased activity toward D-glucose and G4 (Table 3). However, it is unclear if a 96-h incubation is sufficient for the full exchange of the Ca2+ for Zn2+. It remains an intriguing possibility that the enzyme’s 2-3 times enhanced activity in the presence of 0.02 – 0.1 mM Fe2+ or Fe3+ (Mikkelsen et al., 2011) is related to the binding of an iron ion at the position of the calcium ion in the active site.

Figure 5. Stereo image of the binding of a PEG fragment in the substrate-binding site of PQQ-ADH. Amino acids are shown as cyan sticks. The PQQ cofactor is shown as grey sticks, the PEG fragment as yellow sticks and the zinc ion as a brown sphere, metal-ligand bonds are indicated with black dashes. The final σA-weighted 2Fo - Fc map electron density map, in grey, is contoured at 1σ.

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Crystal structure of PQQ-ADH ......

The exchange of the calcium ion by other ions has also been achieved in other quinoproteins. The calcium ion in the Type I QADH from P. aeruginosa could be removed by treatment with a chelating agent and reconstitution by incubation with PQQ and Ca2+ or Sr2+, but not with Mg2+, Mn2+, or Cd2+ (Mutzel, 1991). Likewise, a functional MDH was obtained after reconstitution with Ba2+ or Sr2+ (Adachi et al., 1990, Goodwin et al., 1996a, Goodwin et al., 1996b). Moreover, in the crystal structure of Methylacidiphilum fumariolicum MDH a lanthanide ion was found at the calcium position (Pol et al., 2014). In these other proteins the active site is more compact and the calcium ion is not readily accessible from the solvent. Because of the open, accessible conformation of the active site of PQQ-ADH, the exchange of Ca2+ for Zn2+ appears to be much easier in the latter enzyme.

Substrate binding to PQQ-ADH

The G4-binding experiment revealed positive Fo – Fc electron density near the active site Zn2+ ion and PQQ. The electron density was compatible with a partially occupied PEG550 MME molecule from the crystallization solution, but not with a maltotetraose molecule. In the refined structure the hydroxyl group of the PEG molecule is hydrogen bonded to Asp333 (OD2 atom at 2.5 Å), O5 of PQQ (at 2.3 Å), and Zn2+ at 2.9 Å. Van der Waals interactions with Phe434, Leu578, Tyr476, Leu153, Leu435, and PQQ completes the binding (Fig. 5). The hydroxyl group has exactly the same position as one of the carboxylate oxygen atoms of the tetrahydrofuran-2-carboxylic acid product found in the structure of QH-ADH (Oubrie et al., 2002), after superimposition of the two enzymes. From this, we conclude that the PEG fragment is bound at the expected position for a substrate/product molecule.

Mechanism of oxidation

The PQQ ring structure in native PQQ-ADH is entirely planar, indicating its oxidized orthoquinone (Williams et al., 2005) or reduced hydroquinone PQQH2 form (Oubrie et al., 2002). However, in the G4 structure the PQQ-O4 is out of plane by about 45o (Fig. 5 and S2) and is pulled below the PQQ tricyclic plane towards the Nη of Lys378 and the carboxylate atoms of Asp439. This out-of-plane position of O4 suggests that the C4 atom is sp3 hybridized and bears a hydroxyl group. A similar position of O4 has also been observed in the MDHs from M. extorquens (Ghosh et al., 1995) and M. fumariolicum (Pol et al., 2014). Reduced PQQ is usually protonated at both O4 and O5 (Kay et al., 2006b) or singly protonated at O4 (Kay et al., 2006a, Duine, 1999, Anthony, 1996). Likely, in PQQ- ADH only the first reduction step has occurred with protonation only of the O4 atom, similar to what has been suggested for quinoprotein alcohol dehydrogenase from P.

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aeruginosa (Kay et al., 2006b). Another possibility is that the PQQ is in the radical semiquinone state, which also has the O4 out of the plane of the cofactor (Anthony, 1996). In quinoproteins a hydride transfer mechanism is generally accepted for the oxidation of substrates (Oubrie et al., 1999, Anthony, 1996, Oubrie et al., 2000, Anthony et al., 2003). The mechanism involves proton abstraction from the hydroxyl group of the substrate by a general base catalyst. In M. extorquens MDH and P. aeruginosa QADH the invariant Asp303 (Asp333 in PQQ-ADH) was proposed as the general base (Kay et al., 2006a, Anthony, 1996, Afolabi et al., 2001), while Glu171 was suggested to have this role in Methylophilus W3A1 MDH (Zhang et al., 2007). The residue in PQQ-ADH equivalent to Glu171 is Gln220, which is highly unlikely to function as a general base, and its side chain Oε1 atom is too far (3.7 Å) from the PEG substrate to abstract a proton from its hydroxyl group. Since the invariant Asp is also conserved in PQQ-ADH (Fig. S1), and at hydrogen- bonding distance to the PEG hydroxyl group, we suggest that this residue (Asp333) is the general base in the reaction mechanism of this class of PQQ-dependent dehydrogenases. In the subsequent step of the reaction mechanism (or concomitantly) a hydride ion is abstracted from the hydroxyl-bearing carbon atom, and transferred to the electrophilic C5 carbonyl carbon of the PQQ quinone, upon which PQQ tautomerizes to the hydroquinone PQQH2 (Oubrie et al., 1999, Kay et al., 2006b, Oubrie et al., 2000) and product is released or further converted. The conserved Lys378 (or Arg in some other enzymes) may help in transferring the electron from PQQH2 to an electron acceptor (Yamada et al., 2003). To further characterize the binding mode of substrates in the active site of PQQ-ADH, we carried out docking studies with monosaccharides. These studies indicated that the enzyme can easily accommodate these substrates, both in the pyranose and linear conformation, because of its shallow and solvent exposed active site. These studies also showed that β-D-glucose can be oxidized easier than α-D-glucose because in β-D-glucose the C1 hydrogen atom, which is to be transferred to the PQQ as a hydride ion, has an axial orientation, and faces the C5 atom of the PQQ. Moreover, productive binding of α- cyclodextrin appeared possible, although the C6 carbon atom is probably not close enough for efficient direct hydride transfer, explaining the low activity with this substrate (Mikkelsen et al., 2011).

CONCLUSION

In conclusion, the structure of PQQ-ADH presented here is the first structure of a monomeric Type I quinoprotein alcohol dehydrogenase, distinctly different from other Type I enzymes. The presence of a polyethylene glycol fragment bound in the active site suggests Asp333 as the general base in the catalytic mechanism. The active site is open

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Crystal structure of PQQ-ADH ...... and easily accessible from the solvent and can accommodate linear alcohols, as well as mono- and oligosaccharides of different sizes. Monosaccharides can be bound in both the pyranose and linear conformation. Surprisingly, the enzyme has no vicinal disulfide bond between adjacent cysteine residues, which is observed in other quinone-dependent alcohol dehydrogenases. The absence of this vicinal disulfide may be required to make the active site accessible to substrates of widely different sizes and to create a versatile enzyme.

ACKNOWLEDGEMENTS

We thank Mr. K.H. Kalk for his technical assistance with the data collection in house. We are grateful to the staff scientists at beam line X11 at the EMBL-Hamburg outstation (DESY), Germany, for their support during data collection.

SUPPORTING MATERIAL

Figure S1. Structure–based alignment of PQQ-ADH from Pseudogluconobacter saccharoketogenes IFO 14464, ADHIIG (Toyama et al., 2005) and ADHIIB (Chen et al., 2002) from Pseudomonas putida HK5, QH-ADH from Comamonas testosteroni (Oubrie et al., 2002), QADH from Pseudomonas aeruginosa (Keitel et al., 2000) and MDH from Methylacidiphilum fumariolicum SolV (Pol et al., 2014), Methylobacterium extorquens (Williams et al., 2005), Hyphomicrobium denitrificans (Nojiri et al., 2006) P. denitrificans(Xia et al., 1999) and Methylophilus W3A1 (Li et al., 2011). The structural alignment was made with Promals3D (Pei et al., 2008). The secondary structure elements above the sequence alignment are those obtained from the crystal structure of PQQ-ADH. Identical residues have a red background color and similar residues have a red color. Residues indicated with blue stars below the sequences are PQQ ligands, with purple stars are active site calcium ligands and with grey stars above the sequence have alternate conformations. The vicinal disulfide bridge has a cyan background. Green boxes indicate the tryptophan docking motifs. Disulfide linkages are indicated in green italics below the sequences. The figure was created with ESPript (Gouet et al., 1999).

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Figure S2 Overlay of the structure of PQQ-ADH with calcium bound in the active site in green and with zinc bound in cyan. The calcium ion is shown as a green sphere and the zinc ion as a brown sphere.

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Crystal structure of Endo-Xylogalacturonan Hydrolase from Aspergillus tubingensis

Henriëtte J. Rozeboom1, Gerrit Beldman2, Henk A. Schols2 and Bauke W. Dijkstra1

1 Laboratory of Biophysical Chemistry, Groningen Biomolecular Sciences and Biotechnology Institute, University of Groningen, Nijenborgh 7, 9747 AG Groningen, The Netherlands. 2 Laboratory of Food Chemistry, Wageningen University, Bornse Weilanden 9, 6708 WG Wageningen, The Netherlands

FEBS Journal (2013) 280 (23), 6061-6069

......

Endo-xylogalacturonan hydrolase features a wide-open active site groove to accommodate its xylogalacturonan substrate. No specific xylose binding pockets are present that could hinder sliding movements of the substrate of this processive enzyme. A model of xylosylated tri-galacturonate in the active site explains the enzyme’s preference for hydrolysis of the glycosidic bond linking two β-xylose substituted galacturonic acid residues.

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ABSTRACT

Endo-Xylogalacturonan hydrolase is a member of glycoside hydrolase family 28 (GH28) that hydrolyzes the glycosidic bond between two β-xylose substituted galacturonic acid residues in pectin. Presented here is the X-ray crystal structure of the endo- xylogalacturonan hydrolase from Aspergillus tubingensis (XghA) at 1.75 Å resolution. The high degree of structural conservation in the active site and catalytic apparatus, compared with polygalacturonases, indicates that cleavage of the substrate proceeds in essentially the same way as found for the other GH28 enzymes. Molecular modeling of a xylosylated tri-galacturonate in the active site identified the amino acid residues involved in substrate binding. They border a substrate-binding cleft, that is much wider than in other polygalacturonases, and can accommodate xylosylated substrates. The most extensive interactions appear to occur at subsite +2, in agreement with the enzyme kinetics results, which showed enhanced activity on substrates with a xylose attached to the galacturonic acid bound at subsite +2.

INTRODUCTION

Pectin is a mixture of highly heterogeneous and branched polysaccharides that constitutes a major component of the primary cell wall of plants (Wong, 2008). The predominant use of pectins are in the food and cosmetic industries as gelling and thickening agents, and as stabilizers. Pectins also have positive effects on human health and have multiple biomedical uses (Jayani et al., 2005, Mohnen, 2008). The structure of pectin consists of ‘smooth’ regions of homogalacturonan, and ‘hairy or ramified’ regions of rhamnogalacturonan I, rhamnogalacturonan II, xylogalacturonan (XGA), apiogalacturonan, and polysaccharides comprising mostly neutral sugars (Wong, 2008). The most abundant pectin, homogalacturonan, is a linear homopolymer of α-1,4- linked D-galacturonic acid (GalA) residues, with the carboxylic acid groups methyl- esterified to various degrees up to 70–80%. Xylogalacturonan (XGA) is a polymer of GalA units in which the O-3 atoms of 20-100% of the GalA residues carry a β-(1,3)-linked xylose side chain (Wong, 2008, Coenen et al., 2007, Yapo, 2011). XGA from pea hull (Pisum sativum) contains mainly single terminal xylose residues and a few short 1,2- linked, or sometimes 1,3-linked xylosyl oligosaccharides attached to O-3 (Le Goff et al., 2001). In XGAs from pectin of apple (Malus domestica) and potato (Solanum tuberosum), 1,4-linked xylose disaccharides are observed (Zandleven et al., 2006), while in pectin from soybean (Glycine max) branched XGAs occur with both 1,4 and 1,2 linkages. (Nakamura et al., 2002)

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Crystal structure of XghA ......

Polygalacturonases (PGs) produced by plants are involved in the ripening process to soften and sweeten fruits. In contrast, bacterial and fungal polygalacturonases assist in the degradation process to liberate nutrients on which they can grow. The filamentous fungi of the Aspergillus produce many extracellular hydrolytic enzymes to degrade the pectin backbone (de Vries et al., 2001), and these have been classified in the pectin degrading glycoside hydrolase family 28 (GH28). GH28 includes endo- polygalacturonases, exo-polygalacturonases and rhamnogalacturonan hydrolase (RGase A) (Cantarel et al., 2009). All family GH28 members act with an inverting mechanism (Zandleven et al., 2005). Among the GH28 enzymes, endo-xylogalacturonan hydrolase (XghA, E.C. 3.2.1.-) plays a unique role in the degradation of XGA. XghA is a processive enzyme that cleaves the β- xylose substituted GalA backbone of pectin in an endo-manner, with xylose substitution being a prerequisite (van der Vlugt-Bergmans et al., 2000), although activity has also been observed towards di-galacturonic acid of which only one residue is xylosylated (Zandleven et al., 2006, Zandleven et al., 2005, Beldman et al., 2003). Mature XghA is a 388-residue protein with ~ 22-27% sequence identity to polygalacturonases from other Aspergillus species, but the structural determinants of its typical xylogalacturonase activity have remained obscure. The four active site sequence motifs that are absolutely conserved in GH28 are also present in XghA (Armand et al., 2000, Markovič et al., 2001). The optimal activity of XghA was found between pH 3.0-3.5, and its optimal temperature was ~ 50 oC (Beldman et al., 2003). The enzyme appeared to be stable over a broad pH range (2.5-6.5). Here we present the 1.75 Å crystal structure of endo-xylogalacturonan hydrolase, and compare it to other GH28 family members, revealing that it has a more open active site to accommodate the xylose residues. Modeling of a bound (XGA)3 molecule suggests that the most extensive interactions with xylose occur at subsite +2, in agreement with previous kinetic studies.

RESULTS AND DISCUSSION

Overall structure of endo-xylogalacturonan hydrolase

The crystal structure of endo-xylogalacturonan hydrolase (XghA) was determined to a resolution of 1.75 Å using molecular replacement. Data collection and phasing are summarized in Table 1. The asymmetric unit contains one XghA monomer, one glycerol molecule, seven sulfate ions, and 458 water molecules, and yielded a final R/Rfree of 16.7/18.9 % after refinement. Further details of the refinement are shown in Table 1. The N-terminal signal peptide (18 residues) and the first 16 residues of the mature

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enzyme were not visible in the electron density maps. Thus, the first residue observed in the electron density map is Ala35.

Table 1. Data collection and refinement statistics. Values in parentheses are for the highest resolution shell.

Parameter Value Diffraction data Native Wavelength (Å) 0.9724 Resolution range (Å) 40.7–1.75 Cell dimensions (Å) a ,b , c 76.4, 121.2, 129.7 Number of unique reflections 60896 (8795) Completeness (%) 99.8 (100.0) Overall I/σ (I) 11.6 (3.7)

Rmerge (%) 10.4 (54.2)

Rpim (%) 4.2 (21.4)

R/ Rfree (%) 16.7/18.9 R.m.s. deviations from ideal values Bond lengths (Å) 0.008 Bond angles (°) 1.327 Protein residues 35 - 406 Water molecules 458 Sulfate molecules 7 Glycerol molecule 1 PDB accession ID 4C2L

XghA folds into a right-handed parallel β-helical structure comprising ten complete turns (Fig. 1), which contain three twisted parallel β-sheets (PB1, PB2a, PB2b and PB3; names are as adopted for the PG2 polygalacturonate hydrolase (van Santen et al., 1999). The β- strands in the parallel β-sheets are connected by four turns, T1, T2a, T2b and T3. All T2a and

T2b turns are short, whereas the lengths of the T1 turns in the C-terminal part (from residue 259) and those of the T3 turns in the N-terminal part (until residue 207) are extended. Three short α-helices are observed in the turns, α2 (residues 102-105) in the

T3 turn of coil 1, α3 (residues 136-144) in the T3 turn of coil 2, and α4 (residues 326-

331) in the T1 turn of coil 9. Three incomplete coils are found at the N- and C-terminal ends (indicated as a–c in Fig. 1B). The C-terminus of XghA (residues 392–406) makes a very tight turn and folds back on the protein, forming interactions with PB1 of coils 9, 10 and c (Fig. 2B). This turn is

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Crystal structure of XghA ...... stabilized by a disulfide bridge between Cys389 and Cys400, and by the C-terminal glycine residue, whose carboxylate oxygen atoms form hydrogen bonds with the main- chain nitrogen atoms of Ser320 and Cys321, as well as by many other hydrogen bonds. This seals off the hydrophobic interior of the β-helix at the C-terminus, while the N- terminus is capped by β1, directly followed by α1 (residues 50-60), similar to what has been observed for endo-PG1 (van Pouderoyen et al., 2003).

Figure 1. Crystal structure of the xylogalacturonan hydrolase XghA (N-terminus on the top, C-terminus on the bottom). A. XghA is displayed in cartoon format and colored in a rainbow gradient from blue, to red. The O-linked glycosylation site is found at T38, whereas the N-glycosylation sites are found at N278 and N301. B. Overview of the parallel β-sheets PB1, PB2a, PB2b and PB3, shown in yellow, orange, red and blue, respectively. The six disulfide bridges are shown in green. C. APBS (Adaptive Poisson-Boltzmann Solver) electrostatic surface potential of XghA at pH4, highlighting the prominently positively charged active site cleft. The residues colored in red are negative, the ones colored in blue are positive and the ones in white are neutral. The negatively charged residues in the active site are masked by the surplus of positive amino acids and the low pH. The electrostatic surface has been homogeneously scaled between ±20 kT/e.

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Figure 2. Close-up view of (A) the N- and (B) the C-terminus of XghA.

Six disulfide bridges are present: Cys37-Cys60 (1), Cys85-Cys88 (2), Cys230-Cys247 (3), Cys321-Cys329 (4), Cys363-Cys369 (5) and Cys389-Cys400 (6) (Fig. 1B and Fig.S1). Four of them (bridges 1, 3, 5 and 6) are conserved among fungal endo-PGs and RGase A (Fig. S1). Cys230, of the third disulfide bridge, follows directly the two catalytic aspartates Asp228 and Asp229 in the amino acid sequence, possibly to ensure correct local folding

(Petersen et al., 1997). The fourth disulfide bridge stabilizes the T1 turn of coil 9; one partner, Cys321, precedes the substrate-binding residue Tyr322 (see below), and may be important for stabilizing the position of this tyrosine. In addition, the backbone amide of Cys321 interacts with the carboxylate oxygen atoms of the C-terminal glycine residue (Fig. 2B). A comparison of XghA with true polygalacturonases shows that XghA has the smallest root mean square difference (on Cα positions) with Aspergillus niger PG1 (van Pouderoyen et al., 2003), Colletotrichum lupini PG1 (Bonivento et al., 2008), Aspergillus niger PG2 (van Santen et al., 1999) and Erwinia carotovora polygalacturonase (Pickersgill et al., 1998), all within 1.7 Å.

Glycosylation

The molecular masses observed in the gel filtration chromatography and SDS-PAGE analyses are significantly higher than the 45 kDa predicted from the amino acid sequence. Glycosylation may cause a higher apparent molecular mass in gel filtration experiments, as well as in SDS-PAGE by decreased binding of SDS to the carbohydrate, compared to polypeptides of the same mass (Leach et al., 1980). An N-linked N-acetylglucosamine (GlcNAc) residue was modeled in electron density extending from the Asn278 side chain. Similarly, at Asn301, two GlcNAc residues were visible. Additional sugar residues are probably present at these sites, but their electron densities were not sufficiently clear to model them. Furthermore, an O-glycosylation site

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Crystal structure of XghA ...... was found at Thr38 (Fig. 2A). At this position, the electron density matches that of a covalently bound mannose residue. In fungi, O-glycosylation has a role in modulating the function of secretory proteins by enhancing their stability and solubility. The O- mannosylation at Thr38 probably affords protection from degradation by proteases (Goto, 2007). The disulfide bridge between Cys37 and Cys60 may have a similar function.

Active site

The active site is located in an open groove/cleft on the surface of the protein. At the entrance to the active site, the width of the groove between T3 Ser146 and T1 Gly323 is significantly larger (~ 15.3 Å) than in other GH28 endo-PGs (in which it ranges from 6.4 Å in endo-PG1 to 11.0 Å in endo-PG2 (Fig. 1) (van Pouderoyen et al., 2003, Bonivento et al., 2008), suggesting easy access for xylosyl side chains along the length of pectin polymer (Fig. 1B). The bottom of the groove contains mainly acidic residues (aspartates), whereas the walls of the groove are lined with basic lysine and arginine residues. The electrostatic surface (Fig. 1C) calculated at pH 4 clearly shows a positively charged active-site groove required for interaction with the negatively charged polygalacturonan backbone. From the alignment shown in Fig. S1, Asp207, Asp228 and Asp229 were identified as the catalytic residues of XghA. They are located on T3 turns 4 and 5 at the bottom of the groove. Based on homology with other GH28 enzymes, Asp228 acts as the proton donor (general acid), while either Asp207 and Asp229 (or both) are expected to activate the catalytic water molecule (base) (van Santen et al., 1999). Other absolutely conserved residues in PGs are Asn205 (178 in PG2) and Tyr322 (291 in PG2), which are necessary for substrate binding (Shimizu et al., 2002). His251 (223 in PG2) maintains the proper ionization state of the general acid Asp228 (Armand et al., 2000). Lys286 (258 in PG2) and the cis-configuration of the peptide between Gly256 and Ser257 are necessary for binding of the subsite -1 C6 carboxylate of the substrate (Shimizu et al., 2002). The additional charges provided by the Nη atoms of Lys234 (Asn207 in PG2) and Lys281 (Asn253) (Fig. S1) may be important for the precise positioning of the C6 carboxylates of the xylosylated polygalacturonate rather than for interacting with a neutral xylose residue. The four polygalacturonase sequence motifs, NTD (NTD205-207 in XghA), G/QDD (DDD 227-229 in XghA), G/SHG (SHG 250-253 in XghA), and RIK (GIK 284-286 in XghA) (Markovič et al., 2001), are largely conserved in XghA. The RG substitution in the RIK motif is typical for xylogalacturonases. The loss of the positively charged Arg284 is compensated for by the Nη atom of Lys281. The smaller lysine compared to arginine may allow room for the xylose residue bound to the subsite -1 GalA residue.

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A closer look at the active site showed the presence of two sulfate ions in the electron density map (Fig. 3). The first sulfate ion is bound at subsite -1 between the N and Oγ atoms of Ser257 of the cis-peptide bond, the OH of Tyr322, the Nη atoms of Lys234 and Lys286, and two water molecules. The second sulfate ion, at a distance of 6.4 Å from the first one, is bound at subsite +1 between Nδ of His251, OH of Tyr322, Nη of Lys286 and seven water molecules. This sulfate has an identical position to a sulfate ion observed in PG-1 (van Pouderoyen et al., 2003). Sulfate ions often mimic the positions of saccharide carboxylate groups (van Pouderoyen et al., 2003, van Asselt et al., 1999). Indeed, the two sulfate ions match the positions of carboxylates of the substrate bound to endo-PG1 from Stereum purpereum (Shimizu et al., 2002). Likewise, in Yersinia enterocolitica exopolygalacturonase the carboxylate of the bound GalA at subsite -1 has exactly the same position as the sulfate bound at subsite -1 in XghA (Abbott et al., 2007). From this, we conclude that Tyr322, Lys234, and Lys286 are probably involved in binding the substrate’s carboxylate at subsite -1, and His251, Tyr322, and Lys286 probably interact with the carboxylate of the sugar residue bound at subsite +1.

Figure 3. Close-up view of the active site of XghA. Sulfate ions, binding at the -1 and +1 subsites are shown in ball- and-stick representation. Catalytic residues, D207, D228 and D229, are shown. No interactions occur between the catalytic residues and the sulfate ions. Hydrogen bonds are indicated with dashes.

Substrate modeling and molecular docking

The major difference in the substrates of endo-PGs and XghA is the presence of xylose bound at the O3 of GalA. The conformation of GalA is restricted by the di-axial α-1,4- glycosidic bond and a weak O2···O6’ hydrogen bond (Braccini et al., 1999). In contrast,

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Crystal structure of XghA ...... the xyloses in XGA have more conformational flexibility due to their di-equatorial β-1,3- glycosidic linkage.

From the models of (XGA)2 bound to XghA produced by AutoDock Vina only those were considered that had their galacturonic acid carboxylates at similar positions to the two sulfates in the active site in the crystal structure. Furthermore, candidate models had to have their reducing end (C1) directed toward the C-terminus and their non-reducing end (C4) directed toward the N-terminus of the protein, and both ends had to be accessible from the solvent. As the active site in XghA is open, many XGA models generated by

AutoDock did not interact with the protein. The (XGA)2 model that best fitted the criteria could be extended to the +2 subsite (Fig. 4), but extending this (XGA)3 model to the -2 subsite only gave unacceptable models, probably because of insufficient interactions with the protein. In subsite -1, the carboxylate has interactions with Lys234 and the backbone amide and hydroxyl of Ser257 (cis-peptide). There is enough space to accommodate a xylose residue attached to the O3 of GalA. This xylose residue may interact with the polar Asn174; in the PGs, the equivalent residue is a small hydrophobic residue. At position 175, a Val is present, which is smaller than the Gln conserved in endo-PGs, providing space for the xylose; a Gln at this position would probably clash with the xylose, thereby preventing binding and degradation of the substrate.

Figure 4. Stereoview of (XGA)3 modeled in the active site of XghA. For clarity reasons only amino acids interacting with the substrate are shown at the +1 and –1 subsites. Catalytic residues, D207, D228 and D229, are not shown. Carbons of the GalA backbone are colored green and carbons of the xylose residues are colored cyan.

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In subsite +1, the carboxylate shows interactions with His251 and Lys286. Sufficient space is also available for a xylose branch in this subsite. In contrast, in endo-PGs, the xylose would likely clash with His177 (equivalent to the more flexible Lys204) and with Asn126, which restricts access to the active site because of earlier unwinding of helix α3 in endo-PG. At subsite +2, the carboxylate of GalA interacts with Lys204; the xylose attached to the GalA in this subsite is bound by Thr280, Lys281, Gln317 and Gln319. The scissile glycosidic bond between the -1 and +1 subsites is somewhat strained (φ/ψ =

-22/113) compared to the ideal torsion angle values of polygalacturonan helices (31 helix, φ/ψ = 80/89; 21 helix, φ/ψ = 80/161 (Scavetta et al., 1999) suggesting that interactions with the protein distort its low-energy conformation. In contrast, the glycosidic torsion angles of the β-1,3-bound xylose have more freedom (Table 2), and probably depend on precise interactions of the xylose with the protein. From the modeling it, may be concluded that the most extensive interactions of XghA with xylose exist at the +2 subsite, but no clear pocket is present. Degradation studies with GalA4Xyl3 as a substrate (in which the reducing end galacturonic acid is not xylosylated)

(Zandleven et al., 2005) indicate fast formation of GalAXyl and GalA3Xyl2 products by

XghA. Further conversion of the latter product to GalA2Xyl was much slower. As XghA mainly acts from the non-reducing end on these small substrates, it can be concluded that a xylose residue at the +2 site increases the reaction rate significantly, in accordance with the crystal structure and the modeling study. The inability of XghA to degrade linear oligosaccharides of GalA or minimally β-xylose substituted GalA may be attributed to lack of interaction of the substrate with the subsites -1, +1 and +2.

Table 2. Torsion angles (φ and ψ) of the glycosidic bonds in modeled XghA-(XGA)3 complex structures.

Definitions of the glycosidic bond torsion angles between two (1,4)-linked GalA residues: φ (phi) = (O5i - C1i – O4i+1

– C4i+1) and ψ (psi) = (C1i – O4i+1 – C4i+1 – C5i+1). Definitions of the glycosidic bond torsion angles between GalA and a (1,3)-linked Xyl: φ (phi) = (O5i - C1i – O3i+1 – C3i+1) and ψ (psi) = (C1i – O1i+1 – C3i+1 – C4i+1).

α-1,4 glycosidic bond φ (1,4) ψ (1,4) -1/+1 - 22 113 +1/+2 105 66 β-1,3 glycosidic bond φ (1,3) ψ (1,3) -1 60 84 +1 124 12 +2 74 99

XghA has been reported to be a processive, endo-acting enzyme (Zandleven et al., 2005, Beldman et al., 2003). Indeed, the extended open groove of XghA, where the xylogalacturon polymer binds, is in agreement with this endo-activity. Although an extended substrate-binding groove is a requirement for processivity (Breyer et al.,

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2001), the structural basis for processivity is not obvious, as the substrate is only slightly enclosed by the walls of the groove. However, the enzyme binds the substrate at least at four adjacent sites at least (Zandleven et al., 2005), which may facilitate sliding of the substrate through the groove (Breyer et al., 2001), and thus be a determinant of processivity. In this respect, it is important to note that no obvious binding pockets for the xylose side chains exist that may hinder easy sliding motions.

The crystal structure of XghA constitutes the first endo-xylogalacturonan hydrolase 3D- structure described so far. The structure of XghA, although similar to polygalacturonase structures, reveals a much wider active-site groove and specific xylose-binding interactions at the +2 site. The -1 and +1 subsites also provide extra room for xylose residues. A substrate docking analysis explains the enzyme’s unique substrate specificity for xylosylated polygalacturonan. Further mutational experiments are required for validation of the assigned subsites for XghA binding.

EXPERIMENTAL PROCEDURES

Protein expression and purification

The enzyme xylogalacturonan hydrolase (XghA) from Aspergillus tubingensis was cloned (van der Vlugt-Bergmans et al., 2000) and expressed in Aspergillus niger PlugBug™ (van Dijck, 1999) (kindly provided by DSM Food Specialties Delft, the Netherlands). The crude enzyme preparation was extensively dialyzed against 10 mM Tris buffer, pH 7.5, and filtered over a 0.22 μm filter to remove insoluble debris. XghA was partially purified by anion exchange chromatography on Q-Sepharose HP (GE Healthcare) with 20 mM Tris buffer, pH 7.5, and elution with 300 mM NaCl. The enzyme was further purified by gel filtration using a Superdex75 HR10/30 column (GE Healthcare, Diegem, Belgium), equilibrated with 20 mM Tris buffer, pH 7.5, containing 150 mM NaCl. XghA eluted at a molecular mass of 56 kDa; on SDS-PAGE the apparent molecular mass was ~ 55 kDa. XghA fractions were pooled and concentrated to 4.5 mg mL-1 using an Amicon Ultracel- 30K filter unit (Merck Millipore, Billerica, MA, USA). Dynamic light scattering experiments were performed using a DynaPro MS800TC instrument (Wyatt Technology Corporation, Santa Barbara, CA, USA) at 20 °C. Dynamic light scattering data were processed and analyzed with Dynamics (Wyatt Technology Corporation) software. From the dynamic light scattering experiments an apparent molecular mass of 30 kDa was deduced with a polydispersity of 20%.

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Crystallization, data collection and structure determination

Initial vapour-diffusion crystallization experiments were performed using a Mosquito crystallization robot (Molecular Dimensions Ltd., Newmarket, UK). In a typical experiment, 0.1 μl screening solution was added to 0.1 μl protein solution on a 96-well MRC2 plate (Hampton Research, Aliso Viejo, CA, USA); reservoir wells contained 60 μl screening solution. The screening solutions used for the experiments were Structure Screen (Molecular Dimensions Ltd.), Wizard and Cryo (both from Emerald Biosystems, Bainbridge Island, WA, USA), and JCSG+ (Qiagen Systems, Germantown, MD). XghA crystals were obtained at room temperature from high ammonium sulfate concentrations. Optimization of the crystallization conditions yielded well-diffracting crystals that grew within several weeks to months when 2 µl protein solution (4.7 mg mL-1) was mixed with 2 µl reservoir solution containing 2.9 M ammonium sulfate with 100 mM sodium acetate buffer, pH 4.4, and 15% glycerol. A native dataset to 1.75 Å resolution was collected on beamline ID23-1 at the European Synchrotron Radiation Facility (Grenoble, France) at 100 K. Intensity data were processed using iMOSFLM (Battye et al., 2011) and the CCP4 package (Winn et al., 2011). The space group was I222, with unit cell dimensions of a = 76.4, b = 121.2, c = 129.7 Å. 3 With one monomer of 40 kDa in the asymmetric unit, the VM is 3.8 Å /Da (Matthews, 1968) with a calculated solvent content of 67%. A summary of data collection statistics is given in Table 1. Using the FFAS03 server (Jaroszewski, L., Rychlewski, L., Li, Z., Li, W. & Godzik,A., 2005) and SCWRL (Canutescu et al., 2003), homology models for XghA were generated. Four structures with highest identity were used as templates - A. niger Endo-PG I (PDB code 1NHC; (van Pouderoyen et al., 2003)), A. aculeatus RGase A (PDB code 1RMG; (Petersen et al., 1997)), A. niger Endo-PG II (PDB code 1CZF; (van Santen et al., 1999)), and Colletotrichum lupini Endo-PG1 (PDB code 2IQ7; (Bonivento et al., 2008)) – with sequence identities of 27%, 25%, 22% and 22%, respectively. Molecular replacement was performed using PHASER (McCoy et al., 2007). ARP/wARP (Langer et al., 2008) was used for automatic building and the model was refined using REFMAC5 (Murshudov et al., 1997). COOT (Emsley et al., 2004) was used for manual rebuilding and map inspection. In 2Fo-Fc maps clear electron density was present for N-glycosylation at Asn278 and Asn301, and O-glycosylation at Thr38. The quality of the model was analyzed using MolProbity (Davis et al., 2007); secondary-structure elements were assigned with DSSP (Kabsch et al., 1983). Figures were prepared using PyMOL version 1.2r1 (Schrödinger LLC, New York, NY, USA) (DeLano, 2002) and ESPript (Gouet et al., 1999). Electrostatic profiles were calculated using the APBS (Adaptive Poisson- Boltzmann solver) tool interfaced to PyMOL after a PDB2PQR conversion (Andre-Leroux et al., 2009). Calculations were performed at pH 4 using the AMBER force field. Atomic

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Crystal structure of XghA ...... coordinates and experimental structure factor amplitudes for XghA have been deposited in the RCSB Protein Data Bank under accession code 4C2L.

Substrate modeling and molecular docking

As a starting model for molecular docking, the experimentally observed digalacturonate bound to Yersinia enterocolitica YeGH28 (PDB entry 2UVF, (Abbott et al., 2007)) was used. Xylose residues were added to the C3 atoms of the two galacturonic acids using the

PRODRG server (Schüttelkopf et al., 2004), resulting in the substrate (XGA)2. The modeled substrate was used for automated docking to XghA using AutoDock Vina (Trott et al., 2010) with the aid of the PyMOL plug-in (Seeliger et al., 2010). All solvent molecules were removed from the XghA structure except for the catalytic water molecule. XghA and (XGA)2 were treated as a rigid protein and a flexible molecule, respectively. The grid of the docking simulation was defined by a 22 ⨯ 36 ⨯ 25 Å box centered on the XghA -1 and +1 subsites. The docking simulation was performed using the default parameters. The 20 top-ranked generations based on the predicted binding -1 affinity (kcal.mol ) were analyzed. The best docked (XGA)2 molecule was extended to the +2 subsite by adding a XGA moiety to (XGA)2. The resulting (XGA)3 molecule was used for another docking simulation.

ACKNOWLEDGEMENTS

We thank Marga Herweijer (DSM Food Specialties, Delft, the Netherlands) for her kind preparation of XghA. We are grateful to the staff scientists at beam line ID23-1 at the ESRF, Grenoble, France, for their support during data collection.

SUPPORTING MATERIAL

Figure S1. Structure–based alignment of GH28 enzymes. Structure–based alignment of XghA from Aspergillus tubingensis, endo-polygalacturonase I from Aspergillus niger (PG1), endo-polygalacturonase II from Aspergillus niger (PG2), polygalacturonase from Aspergillus aculeatus (PGA), and rhamnogalacturonase A from Aspergillus aculeatus (RGase). The structural alignment was made with Dali (Holm et al., 2000). The secondary structure elements above the sequence alignment are those obtained from the crystal structure of XghA. Identical residues have a red background color, similar residues have a red color. Residues in the –1 subsite are shown with a purple background, and the catalytic residues have a cyan background. Residues marked with a * (asterisk) have double conformations. Disulfide linkages are indicated in green italics below the sequences. The figure was created with ESPript (Gouet et al., 1999).

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Chapter 6

Crystal structures of two Bacillus carboxylesterases with different enantioselectivities

Henriëtte J. Rozebooma, Luis F. Godinhob*, Marco Nardinia#, Wim J. Quaxb and Bauke W. Dijkstraa aLaboratory of Biophysical Chemistry, Centre of Life Sciences, University of Groningen, Nijenborgh 7, 9747 AG Groningen, The Netherlands. b Department of Pharmaceutical Biology, Groningen Research Institute of Pharmacy, University of Groningen, Antonius Deusinglaan 1, 9713AV Groningen, The Netherlands. # Present address: Department of Biosciences, Via Celoria 26, University of Milano, 20133 Milano, Italy. *Present address: Department of Biology, Center of Cell Biology, University of Aveiro, Campus de Santiago, 3810-193 Aveiro, Portugal

Biochimica et Biophysica Acta (2014) 1844, 567-575

......

Crystal structures of carboxylesterases NP and CesB have been determined. Their α/β hydrolase fold resembles that of C-C bond cleaving MCP hydrolases. Their different enantioselectivities result from only a few amino acid substitutions.

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ABSTRACT

Naproxen esterase (NP) from Bacillus subtilis Thai I-8 is a carboxylesterase that catalyzes the enantioselective hydrolysis of naproxenmethylester to produce S-naproxen (E > 200). It is a homolog of CesA (98% sequence identity) and CesB (64% identity), both produced by B. subtilis strain 168. CesB can be used for the enantioselective hydrolysis of 1,2-O-isopropylideneglycerol (solketal) esters (E > 200 for IPG-caprylate). Crystal structures of NP and CesB, determined to a resolution of 1.75 Å and 2.04 Å, respectively, showed that both proteins have a canonical α/β hydrolase fold with an extra N-terminal helix stabilizing the cap subdomain. The active site in both enzymes is located in a deep hydrophobic groove and includes the catalytic triad residues Ser130, His274, and Glu245. A product analog, presumably 2-(2-hydroxyethoxy)acetic acid, was bound in the NP active site. The enzymes have different enantioselectivities, which previously were shown to result from only a few amino acid substitutions in the cap domain. Modeling of a substrate in the active site of NP allowed explaining the different enantioselectivity. In addition, Ala156 may be a determinant of enantioselectivities as well, since its side chain appears to interfere with the binding of certain R-enantiomers in the active site of NP. However, the exchange route for substrate and product between the active site and the solvent is not obvious from the structures. Flexibility of the cap domain might facilitate such exchange. Interestingly, both carboxylesterases show higher structural similarity to meta-cleavage compound (MCP) hydrolases than to other α/β hydrolase fold esterases.

INTRODUCTION

Nature has evolved a diverse range of esterases with high regio- and stereoselectivity to convert a broad variety of chemical compounds in various metabolic processes (Bornscheuer, 2002a). Because of their specificity and selectivity they are frequently used in biotechnological applications, such as in the food industry, for the production of perfumes, and for the synthesis of pharmaceuticals (Jaeger et al., 2002). Carboxylesterases (carboxylic acid ester hydrolases, E.C. 3.1.1.1.) are a subset of esterases that hydrolyze small, water-soluble ester containing molecules resulting in the formation of an alcohol and a carboxylic acid. The microorganism B. subtilis strain Thai I- 8 produces an enantiospecific carboxylesterase, called naproxen esterase or carboxylesterase NP (NP), which can be applied as a biocatalyst for the enantioselective resolution of racemic 2-arylpropionates, such as the important non-steroidal anti- inflammatory drugs naproxen, ibuprofen, indomethacin, and nabumetone. NP has a product ee of 96% based on the substrate S-naproxenmethylester (Fig. 1) and 99% on S- naproxenethylester (E > 200) (Quax et al., 1994, Steenkamp et al., 2008). In contrast, the

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Crystal structures of two Bacillus carboxylesterases ...... enzyme’s enantioselectivity towards 1,2-O-isopropylideneglycerol esters (IPG esters) (Fig. 1) is not sufficient for industrial processes (E values of ~ 1.2 for C4 and 1.3 for C8 IPG esters, respectively) (Dröge et al., 2001). (S)-IPG (D-(+)-solketal) is an important building block for the synthesis of the biologically active forms of several well-known pharmaceuticals and endogenous compounds such as β-blockers, prostaglandins, and leukotrienes (Dröge et al., 2001, Godinho et al., 2012).

Figure 1. Two-dimensional depiction of the reaction catalyzed by the carboxylesterases. The * indicates the chiral carbon atom

B. subtilis strain 168 contains two homologs of NP called CesA (98% identical to NP; also known as carboxylesterase NA) and CesB (64% identical to NP; also known as YbfK). Unlike NP, CesB does convert (S)-IPG-caprylate ester with high enantioselectivity to (S)- IPG (ee of 99.9%, E > 200), but not (R)-IPG-caprylate ester (Fig. 1) (Dröge et al., 2001). No marked differences in activity and selectivity were observed between NP and CesA (Dröge et al., 2005). All carboxylesterases of which 3D-structures have been elucidated to date share high structural similarity; all contain the characteristic α/β hydrolase fold (see the ESTHER database) (Lenfant et al., 2013). This fold is composed of a central β-sheet flanked on both sides by α-helices and serves as a stable protein core where, during evolution, amino acid substitutions, loop insertions or deletions have led to enzymes with diverse catalytic functions (Ollis et al., 1992, Nardini et al., 1999a, Carr et al., 2009, Holmquist, 2000, Kourist et al., 2010). Members of the α/β hydrolase fold have a nucleophile-His-acid catalytic triad with a sequence order of nucleophile, acid, histidine (Heikinheimo et al., 1999). In carboxylesterases the nucleophile is a serine residue, while the acid can be an aspartate or a glutamate residue. The serine residue is the central residue of the conserved pentapeptide sequence motif Gly-X1-Ser-X2-Gly (Bornscheuer, 2002b) which defines a

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‘nucleophilic elbow’ characterized by a very sharp turn between strand β5 and the following α-helix. In the discussed carboxylesterases this motif is Gly-Leu-Ser-Leu-Gly. The catalytic mechanism proposed for carboxylesterases starts with nucleophilic attack by the serine hydroxyl group on the substrate carbonyl carbon atom of the scissile bond. The serine hydroxyl group is activated by the catalytic histidine/acid pair, which takes up the proton from the Ser OH group. A transient tetrahedral intermediate is formed, which is stabilized by two peptide nitrogen atoms, usually from residues on strands β5 and β3, forming the so-called “”. The proton is then transferred from the histidine to the leaving alcohol group, while the acid group of the substrate becomes covalently bound to the serine forming the covalent intermediate. Next, the histidine activates a water molecule, which hydrolyzes the covalent intermediate via nucleophilic attack on the carbonyl carbon of the intermediate. After the hydrolysis, the histidine donates its proton back to the serine, and the acyl component of the substrate is released (Holmquist, 2000, Jaeger et al., 1999). Although the catalytic machinery of α/β hydrolases is very similar, the means by which they bind substrates varies from protein to protein. In most cases, the “cap” or “lid” domain, which shields the catalytic triad, contributes to substrate binding (Carr et al., 2009). This domain is usually inserted at the C-terminal ends of strands 4, 6, 7, or 8, and may differ considerably in size in the various enzymes (Carr et al., 2009). Here, we report the crystal structure of NP at 1.75 Å resolution with a product analog in the active site, as well as the crystal structure of CesB at 2.0 Å resolution. Surprisingly, the NP and CesB structures resemble more those of the C-C bond cleaving (MCP) hydrolases than the α/β hydrolase fold esterases. Modeling of a substrate in the active site of NP allowed to defining the residues that determine their different enantiopecificities, in agreement with previous site-directed mutagenesis results (Godinho et al., 2012).

MATERIALS AND METHODS

Expression, purification, crystallization and structure determination of NP

The nap gene coding for NP from Bacillus subtilis Thai I-8 (CBS 679.85) was cloned into B. subtilis I-85. Cells were grown in 2 ⨯ YT medium and harvested by centrifugation. After treatment of the cells with lysozyme and DNAse, the soluble fraction was harvested by centrifugation. Proteins were precipitated with 60% ammonium sulfate and redissolved in 0.02 M MOPS buffer, pH 7.5, and ultrafiltrated with an Amicon YM filter. The resulting solution was applied to an analytical HPLC-SEC column (TSK 2000 SW, 2 times 300 ⨯ 7.5 mm), and the protein was eluted with 0.01 M MES (2-(N-

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Crystal structures of two Bacillus carboxylesterases ...... morpholino)ethane sulfonic acid), pH 5.6, and 0.1 M NaCl with a flow rate of 1 ml/min. The purified enzyme was lyophilized (Bertola et al., 1987). Crystallization of naproxen esterase was performed as described earlier (van der Laan et al., 1993). In short, hexagonal plate-like crystals were grown by liquid-liquid diffusion by filling glass capillaries with 5–10 μl of 80% (w/v) PEG 6000 in 0.1 M Tris-HCl, pH 8.0, and closing them on one side. The equivalent amount of the protein solution (10 mg/ml) in 0.1 M Tris-HCl was added on top of the PEG column and then the capillaries were closed. The temperature was gradually increased over a four-week period from 4 oC to room temperature. Crystals were mounted in a cryoloop and flash-cooled, without adding any cryoprotectant, prior to data collection. Data were collected on the BW7B beamline of the EMBL outstation at DESY (Hamburg, Germany) at 100 K. Intensity data were processed using DENZO and SCALEPACK (Otwinowski et al., 1997). The crystal diffracted to 1.75 Å and belonged to the trigonal space group P3221 (Table 1), with one monomer of 33.8 kDa in the asymmetric unit (corresponding to a calculated VM of 2.1 Å3/Da (Matthews, 1968), and an estimated solvent content of 40%). The crystal structure of NP was solved by molecular replacement with PHASER (McCoy et al., 2007), using a search model composed of three homologous proteins, which were suggested by the Fold and Function Assignment System (FFAS) server (Jaroszewski, L., Rychlewski, L., Li, Z., Li, W. & Godzik,A., 2005). The search model was composed of the structures of Pseudomonas putida esterase (PDB ID: 1ZOI) (Elmi et al., 2005), the meta- cleavage product hydrolase (CumD) from Pseudomonas fluorescens IP01 (PDB ID: 1IUN) (Fushinobu et al., 2005), and the 2-hydroxy-6-oxo-6-phenylhexa-2,4-dienoate hydrolase (BphD) from Rhodococcus sp. strain RHA1 (PDB ID: 1C4X) (Nandhagopal et al., 1997), superimposed on each other. Several cycles of manual re-building with Coot (Emsley et al., 2010), automatic model building with ARP/wARP (Langer et al., 2008), and refinement with REFMAC5 (Winn et al., 2011) were performed in order to improve and to expand the original model of 125 amino acid residues distributed in 6 β-strands and 5 α-helices. Subsequently, a strategy to avoid model bias was applied, employing density modification, solvent flattening, statistical phase improvement, and automated model building performed in cycles using RESOLVE (Terwilliger, 2003). The best partial model generated by RESOLVE contained 180 amino acid residues with additional interpretable electron density, suitable for manual building. The model was completed by a few cycles with ARP/wARP and was further refined with REFMAC5 using TLS refinement (Painter et al., 2006). Water molecules were automatically added by Coot, and manually checked. A 2-(2- hydroxyethoxy)acetic acid molecule was included at the last stage of refinement in the proximity of the nucleophile Ser130. The final model consists of 283 amino acid residues (9-293), 220 water molecules and one 2-(2-hydroxyethoxy)acetic acid molecule and has

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a R factor of 22.4 % and Rfree of 26.4 % (Table 1). The first 8 and last 7 residues are not visible in the electron density and therefore not included in the final model. The stereochemical quality of the model was assessed with MolProbity (Chen et al., 2010) Residue Ser175 has phi/psi angles in the disallowed region of the Ramachandran plot. This strained conformation is due to crystal contacts.

Table 1.Data collection and refinement statistics Values in parentheses are for the highest resolution shell

Data collection NP CesB (YbfK)

Spacegroup P3221 P2 Cell dimensions a, b, c (Å) 47.0, 47.0, 212.5 82.1 , 44.0 , 108.7 αResolution, β, γ (o) range (Å) 90.0,40.0 -90.0,1.75 120.0 90.0,46.0 -91.9, 2.04 90.0 Number of measurements 134364 154210 Number of unique reflections 28534 47148

Rmerge†(%) 5.7 (20.4) 13.3 (39.4) Completeness (%) 99.4 (100.0) 92.8 (76.1) Overall I/σ (I) 21.4 (5.4) 7.1 (2.9) Refinement ¶ Rcryst / Rfree (%) 22.7 / 26.7 20.4 / 25.0 Geometry R.m.s. deviation bonds (Å) 0.008 0.008 R.m.s. deviation angles (o) 1.2 1.2 Contents of asymmetric unit No. of protein atoms 2290 4500 No. ligand atoms 1 2-(2-hydroxyethoxy)acetic acid 4 sodium ions No. water atoms 220 461 Av. main chain B-factor 33.9 17.6 / 20.0 Av. ligand B-factor 49.5 - Av. Water B-factor 40.3 26.1 Ramachandran plot Favored regions (%) 96.8 96.5 Outliers (%) 0.4 0.0 Molprobity Clashscore / percentile 7.1 / 88th 5.2 / 97th Molprobity score / percentile 1.58 / 89th 1.51 / 97th PDB ID 4CCW 4CCY

† Rmerge= ƩhklƩi | Ii(hkl) - 〈I(hkl)〉 | ∕ Ʃhkl Ʃi Ii(hkl). ¶ Rcryst=Ʃhkl| ǀFobsǀ - ǀFcalcǀ| ∕ ƩhklǀFobsǀ, where Fobs is the observed structure factor and Fcalc is the calculated structure factor. Rfree is the same as Rcryst, except that it was calculated using 5% of the data that were not included in any refinement calculations.

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Modeling of the enzyme-substrate complex

Modeling of an enzyme-substrate complex was performed with the program Coot. The general position of the tetrahedral intermediate was modeled by superposing the catalytic triad of NP with the catalytic triad of carboxylesterase Est30 from Geobacillus stearothermophilus in complex with a covalently bound tetrahedral reaction intermediate (propyl actetate) (PDB ID: 1TQH) (Liu et al., 2004). Est30 has 16% overall sequence identity to NP (Table S1). Atomic coordinates and structure factor amplitudes have been deposited with the RCSB Protein Data Bank with accession code 4CCW for carboxylesterase NP.

Expression, purification, crystallization and structure determination of CesB

The CesB gene was amplified from the pMAybfk vector (Dröge et al., 2005) using two synthetic primers. The forward primer (5’ – GGGAGGGGCATT CCATGG TACAAGATTC – 3’) contained an NcoI restriction site (in bold) and the reverse primer (5’ – CTTTTCATG AAGCTT TTCCTTACTATTTTATC – 3’) contained a HindIII restriction site (in bold). The resulting PCR product and the pBAD/Myc-HisA vector were digested with NcoI and HindIII restriction enzymes, purified, and ligated using T4 DNA . The ligation mixture was used to transform competent Escherichia. coli TOP10 cells. To verify the presence of the insert, transformants were selected from LB/Amp plates, and analyzed by colony PCR. Positive clones were grown overnight on LB/Amp medium and the plasmid DNA was isolated for sequence analysis. The vector pBAD/Myc-HisA was used for the expression of wild type CesB. E. coli TOP10 cells containing the appropriate expression plasmid were inoculated in 2xLB/Amp o medium (5 ml) and grown overnight at 37 C. Next day the A600 was determined for the overnight cultures and a sufficient quantity of the culture was used to inoculate 2 ⨯ o LB/Amp medium (1 L) to an initial A600 of 0.05. Cultures were grown at 37 C till A600 ≥ 0.5, when the cultures were induced with arabinose (0.02%, w/v) and the temperature was changed to 20 oC. After 16 hours cells were harvested and resuspended in 3 mL/g of wet cells in extraction buffer (10 mM Tris-HCL, pH 8, 1 mM EDTA). Cells were disrupted by sonication, and extracts were clarified by centrifugation (50 min, 40,000g). Purification was performed by anion exchange chromatography using 3 HitrapQ columns (1.6 ⨯ 2.5 cm) with 10 mM Tris-HCl, pH 8.0, and 1 mM EDTA as Buffer A, and Buffer A containing 1.0 M NaCl as Buffer B. Further purification was achieved by phenylsepharose HP chromatography with 10 mm phosphate buffer, pH 8.0, containing 0.5 M ammonium sulfate as Buffer A, and Buffer B containing 10 mm phosphate buffer pH 8.0 (Dröge et al., 2001). In addition, gel filtration using a Hiload Superdex 200 column (GE Healthcare)

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was used to determine the oligomeric state of CesB in solution (monomer) and for buffer exchange (50 mM Tris HCL, 10% (v/v) glycerol, pH 8.0). CesB crystals were obtained from sitting-drop experiments with 20 % polyethylene glycol 3350 and 0.2 M sodium fluoride in 100 mM Bis-Tris propane buffer (pH 6.0–7.0) as precipitant, using drops of 0.1 μl of protein solution (5.2 mg ml-1 in 20 mM Tris buffer, pH 7.5, 100 mM NaCl) and 0.1 μl of reservoir solution set up by the Mosquito crystallization robot. Crystals grew in space group P2 (Table 1) with two monomers of 3 35.0 kDa in the asymmetric unit (corresponding to a calculated VM of 2.8 Å /Da (Matthews, 1968), and an estimated solvent content of 56%). Before data collection, crystals were soaked for 15 s in a cryoprotectant solution, consisting of reservoir solution supplemented with 15 % glycerol, directly followed by flash cooling. X-ray data were collected in house at 110 K with a MarDTB Goniostat System using Cu-Kα radiation from a Bruker MicrostarH rotating-anode generator equipped with HeliosMX mirrors. Intensity data were processed using iMosflm (Battye et al., 2011) and scaled using SCALA from the CCP4 suite (Winn et al., 2011). The structure of CesB was solved by molecular replacement with PHASER (McCoy et al., 2007) using the NP structure as a search model. The phases were improved with the ARP/wARP (Langer et al., 2008) procedure combined with manual model building with Coot (Emsley et al., 2010). TLS refinement (Painter et al., 2006) was performed with REFMAC5 (Winn et al., 2011) and resulted in a final model comprising 2 protein molecules of 285 residues each in the asymmetric unit, 2 sodium ions and 460 water molecules, with a final R factor of 20.4 % and Rfree of 25.0 % (Table 1). No electron density was visible for the first 10 residues and the C-terminal residues of both molecules and these residues were therefore not included in the final model. The stereochemical quality of the model was assessed with MolProbity (Chen et al., 2010). The sodium ions were assigned by the STAN server (Nayal et al., 1996). Atomic coordinates and structure factor amplitudes of carboxylesterase CesB have been deposited with the RCSB Protein Data Bank with accession code 4CCY.

RESULTS AND DISCUSSION

Overall structure

The structures of B. subtilis Thai I-8 carboxylesterase NP and B. subtilis 168 CesB were solved by molecular replacement and refined to 1.75 Å resolution (space group P3221) and 2.04 Å resolution (space group P2), respectively. Data collection and refinement statistics are summarized in Table 1. The asymmetric unit of the NP crystal contains one protein molecule of 283 amino acid residues whereas the asymmetric unit of the CesB crystal contains two protein molecules of 285 residues each. Both carboxylesterases

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Crystal structures of two Bacillus carboxylesterases ...... have a globular shape with dimensions of ~54 ⨯ 51 ⨯ 43 Å3 and share a superimposable tertiary structure with a root mean square deviation (RMSD) value of 0.94 Å for 273 Cα pairs, as expected from their high sequence identity (64%). Like most of the members of the α/ hydrolase fold family, the NP and CesB structures consist of two domains: a central α/ “core” domain, containing the catalytic triad, and an α-helical “cap” region on top of it, responsible for substrate recognition/binding (Fig. 2). It should be noted that NP is 98.0% identical in sequence to B. subtilis strain 168 CesA, differing only in 6 amino acids (L101V, K181T, T188N, Q202K, K217N, A282T) located on the surface of the protein and not influencing the specific activity of the proteins (Dröge et al., 2005). Recently a putative hydrolase (2632844) from B. subtilis strain 168 was deposited with the Protein Data Bank (PDB ID: 2R11) (http://dx.doi.org/10.2210/pdb2r11/pdb) as part of a structural genomics project (Joint Center for Structural Genomics). This enzyme has 100% sequence identity to B. subtilis CesA. As expected, the crystal structures of NP and CesA are very similar, having a RMSD value of 0.55 Å over 282 Cα pairs. CesB is also highly similar to these two enzymes, and therefore the structural description will focus on NP. Specific differences among the structures will be eventually highlighted. The NP core domain comprises residues 30-157 and 224-300, and it consists of a central eight-stranded, predominantly parallel β-sheet flanked by one 310-helix (η2) and three α- helices on one side (α2, α10 and α11) and by four α-helices (α3, α4, α8 and α9) and one

310-helix (η3) on the other side (Figs. 2a, S1 and S2). The core domain has two 310-helices (η2 and η3), a small β-strand (β4’) and one small α-helix (α10) extra compared to the ‘canonical’ α/β hydrolase fold (Ollis et al., 1992, Nardini et al., 1999a, Carr et al., 2009). Additionally, in CesB a small antiparallel β-sheet is formed with β6’ and β6” (Figs. 2B, S2). Furthermore, CesA misses η1 and η4 but has a small β-strand β6’. The cap domain comprises the N-terminal residues 1-29 and residues 158-223 (inserted between strand β6 and helix α8) and consists of four α-helices (α1, α5, α6 and α7) and one 310-helix (η4) (Figs. 2a, S2). The η4 310-helix is located before helix α7 in NP, while it is after α7 in CesB (Fig. 2B). In this class of esterases the most striking feature of the cap domain is the presence of the N-terminal helix α1, which is situated almost parallel to helix α7 and ends as a 310-helix (η1). Interactions between α1 and α7 are mainly hydrophobic. An N-terminal α-helical extension is rare for α/β hydrolase fold proteins, being observed only in the hormone-sensitive lipase subfamily of enzymes such as brefeldin A esterase from B. subtilis (PDB ID: 1JKM) (Wei et al., 1999), Est2 from Alicyclobacillus acidocaldarius (PDB ID: 1EVQ) (De Simone et al., 2000, Mandrich et al., 2009), AFEST, a novel hyper-thermophilic carboxylesterase from the archaeon Archaeoglobus fulgidus (PDB ID: 1JJI) (De Simone et al., 2001) and acetyl esterase HerE from Rhodococcus sp. strain H1 (PDB ID: 1LZL) (Zhu et al., 2003). In these four enzymes the cap domain is built up of two N-terminal α-helices and two α-helices from the insert

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located after strand β6. Kinetic data of Est2 indicate that the first 35 residues of the protein play a role in the conformational stability of the protein and have a regulatory role (Mandrich et al., 2009, Mandrich et al., 2005). It is likely that the N-terminal α-helix in the carboxylesterases has a similar function. A DALI search (Holm et al., 2000) with the α/β hydrolase core domain did come up with many structural neighbors (Table S1). In contrast, for the isolated cap domain no neighbors were found, which makes these carboxylesterases unique in the family of esterases.

Figure 2. Structures of (A) NP from Bacillus subtilis Thai I-8, and (B) CesB from Bacillus subtilis 168. The proteins are shown as cartoons with helices of the core domains colored red/cyan, β-strands yellow/magenta, and the helices of the cap domains slate blue/salmon, respectively. The catalytic triads, S130, H274 and Glu245, for both esterases, are shown as sticks. N and C mark the amino and carboxylate termini of the proteins.

The cap domain is also the protein region where most of the sequence variations are present between NP and CesB (Fig. S1). The most disparate part is at amino acids 209- 218 where differences in amino acid sequence have direct structural implications, such as the presence of the 310-helix η4 (residues 210-212) in CesB but not in NP, or the narrower active site entrance in CesB (Fig. 3). Variations in sequence are also observed in the active sites of NP and CesB (Godinho et al., 2012), explaining the different substrate specificities (see below).

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Figure 3. Entrances to the active sites of NP (A) and CesB (B). (C), structure of NP showing the absence of a back-access channel, blocked by Phe66, in blue and the residues with the largest conformational differences between the carboxylesterases in red. (D), surface plot of NP showing one possible route to and from the active site.

Sodium ion binding to CesB

Two sodium ions, originating from the crystallization solution, are bound to each of the two CesB monomers present in the asymmetric unit (Fig. 4). The first sodium ion (Na1) (Fig. 4A) is bound in a pocket lined by residues 244 to 250 (loop between strand β7 and helix α9). It is octahedrally coordinated by Oε1 of Gln250 (2.34 Å), the carbonyl oxygen atom of Glu245 (2.31 Å), the carbonyl oxygen atom of Tyr248 (2.69 Å), and three water molecules (2.59 Å, 2.53 Å, and 2.46 Å). Interestingly, Glu245 is the acid member of the catalytic triad. There is no sodium ion bound in NP or CesA at this position, where Gln250 is substituted for a Pro. The other sodium ion (Na2) (Fig. 4B) is located 15 Å away from Na1; it has also an octahedral geometry, being coordinated by Thr187 Oγ1 (2.48 Å), the carbonyl oxygen atom of Gly188 (2.32 Å), the carbonyl oxygen atom of Tyr191 (2.45 Å), and three water molecules (2.33 Å, 2.44 Å, and 2.33 Å). The coordinating residues are part of the loop connecting helices α6 and α7 of the cap domain. At this location no sodium ion is bound in NP or CesA; the structures in this region are the most divergent. The binding of sodium could be a result of the purification and crystallization procedures.

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Figure 4. Sodium ions Na1 (A) and Na2 (B) binding to CesB. CesB is depicted as a cartoon in grey. Selected amino acids are shown as sticks colored according to atom type. The catalytic triad is colored in green. Black dashed lines indicate interactions of the sodium ions with the amino acids and water molecules.

The active site

The active site of NP is located at the interface between the core and the cap domains in a ~13 Å deep hydrophobic gorge. Its entrance is lined by residues from helices α5, α6, and α7 and from a loop located after helix α7, all belonging to the cap domain. The active site nucleophile Ser130 is located at the bottom of the gorge, protected from the bulk solvent, on the nucleophilic elbow between strand β5 and helix α4 as part of the conserved sequence motif Gly-Leu-Ser-Leu-Gly. The main chain conformation of this serine is somewhat strained (φ = 53o, ψ = -125o), as more often observed in α/β-hydrolases, but still in the allowed region of the Ramachandran plot. Its conformation is stabilized by a network of hydrogen bonds of the Ser130 carbonyl oxygen atom to the backbone amides of Gly132 (3.1 Å), Gly133 (2.9 Å), and Ala156 (3.0 Å). The Ser130 backbone amide is at hydrogen bonding distance to the carbonyl oxygen atoms of Leu153 (3.0 Å) and Ser154 (3.1 Å). Furthermore, in NP the Ser130 Oγ makes a hydrogen bond to the backbone amide of Leu131, which induces an unfavorable conformation of the nucleophile for the reaction to occur (Fig. 5A). On the contrary, in both CesB monomers the Ser130 Oγ is hydrogen bonded to the Nε2 atom of the catalytic His274 (2.8 Å) (Fig. 5B). This observation suggests the presence of two populations existing for the Ser130 Oγ, one making a hydrogen bond to the backbone amide of Leu131 (representing the so called o “resting” state, with a χ1 of 153 ), another making a hydrogen bond to the His274 side o chain (the “active” state, with a χ1 of -159 ). Indeed, the catalytic His274 in α/ hydrolases is supposed to act as a base, deprotonating the serine to generate a very nucleophilic alkoxide (-O-) group. Interestingly, in CesA, where four independent molecules are present in the asymmetric unit, both populations are present, each associated with two monomers. The resting state found in NP is also reminiscent of that observed for the active site cysteine of dienelactone hydrolase (Carr et al., 2009). Another example is given by B. subtilis lipase, although the resting state of the catalytic

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o serine has a different conformation with a χ1 of 52 (Kawasaki et al., 2002). These different conformations of the catalytic serine side chain support the notion that different orientations of its hydroxyl group are a necessary requirement for the enzymatic activity (Kawasaki et al., 2002). Regarding the third member of the catalytic triad, Glu245 is hydrogen bonded to His274, orienting its imidazole ring such that it interacts with Ser130. Having a glutamate residue as the third member of the catalytic triad is quite atypical; in other α/β- hydrolases and serine esterases normally an aspartate residue is observed. A glutamate as member of the catalytic triad has been found in the family of large esterases (LES, with molecular weights above 55 kDa, subfamily VII (Arpigny et al., 1999) like butyrylcholinesterases, acetylcholinesterases, human carboxylesterase and fungal lipases. Also Est713 (Bourne et al., 2000), a smaller esterase of 35.5 kDa has a glutamate as the third catalytic residue but it is positioned at the end of β6, whereas in the carboxylesterases the glutamate is located at the end of β7, the usual position for the catalytic acid residue in the α/β-hydrolase fold enzymes. Finally, in NP the amide NH groups of Ala64 (Gly64 in CesB) and Leu131 form the oxyanion-binding site that stabilizes the tetrahedral transition-state intermediates (Scheme 1).

Figure 5. View of the active sites, catalytic triads and oxyanion-binding sites of NP (A) and CesB (B). Selected amino acids are shown as sticks colored according to atom type. Amino acids in CesB that differ from the equivalent ones in NP are colored in salmon. The amino acids of the catalytic triad are colored in green. The main chain atoms of Leu131 and Ala/Gly64 are included to mark the oxyanion-binding site. The product, 2-(2- hydroxyethoxy)acetic acid, is colored in cyan. The omit map is shown as a grey mesh and contoured at 1σ.

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Scheme 1. Proposed mechanism of esterase-catalyzed hydrolysis of naproxen methyl ester. The chiral carbon in the substrate is marked with an * (asterisk). The figure has been adapted from Jaeger et al. (Jaeger et al., 1999).

Product in the active site of carboxylesterase NP

An unexpected residual electron density was observed in the active site of NP, near the side chain of Ser130 (Fig. 5A). This electron density is probably indicative of binding (part of) a chain of acidified PEG from the crystallization solution. The crystal was grown from 40% PEG and was stored for several years in this solution before data were collected. It is known that polyethylene glycol solutions get oxidized by air after prolonged storage (Ray et al., 1985). We modeled it as 2-(2-hydroxyethoxy)acetic acid. One of the carboxylate oxygen atoms is weakly hydrogen bonded to Ser130 Oγ (3.8 Å), and His274 Nε2 (3.3 Å), and a water molecule (3.4 Å). The other carboxylate oxygen is hydrogen bonded to the carbonyl group of Ala64 (2.9 Å), of which residue the amide nitrogen atom is involved in the oxyanion binding site (Fig. 5), but no interactions are present between the product analog and the oxyanion binding site amides. Several van der Waals interactions are present instead between the bound molecule and Ala64, Leu65, Phe166, Ala170 and Phe182 at the entrance of the active site. In CesB only water molecules were found in the active site, but in CesA an ethyleneglycol molecule and a triethyleneglycol molecule were modeled in two of the four monomers in

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Crystal structures of two Bacillus carboxylesterases ...... the asymmetric unit. The ethyleneglycol molecule in monomer B (active state) resides in about the same location as the product analog in NP; the triethyleneglycol molecule in monomer C (resting state) is also located at the entrance to the active site but bound via van der Waals interactions to Leu131 and Pro222. This shows the wide-open entrance to the active site of these carboxylesterases.

Structural differences in the NP and CesB active sites and enantioselectivity properties

While (R,S)-ethoxyethyl-[2-(6-methoxy-2-naphtyl)]propionate is enantioselectively hydrolyzed by NP to yield optically active (S)-naproxen, the enzyme hydrolyzes IPG- esters without enantioselectivity (Dröge et al., 2005). This lack of enantioselectivity towards IPG-substrates is mainly due to two residues, Phe166 and Phe182, in the active site as demonstrated by replacing them with the equivalent CesB residues Val166 and Tyr182 (Godinho et al., 2012). In contrast, CesB hydrolyzes 1,2-O-isopropylideneglycerol esters to (S)-IPG with 99.9% enantioselectivity. Thus both esterases are highly selective, but NP is enantioselective towards the acyl/aryl part of substrates, while CesB specifically recognizes the chirality of the alcohol part, with some additional enantioselectivity towards the acyl/aryl part. Since NP (and its homolog CesA) have a high enantiospecificity for S-naproxenethylester (99%) (Dröge et al., 2005), we tried to model the S- and R-naproxenester substrates in the active site of NP (Fig. 6). For the modeling, several interactions were taken into account. For productive binding, the substrate’s ester carbonyl oxygen atom needs to make hydrogen bonds to the two NH-groups of the oxyanion hole (Ala64 in NP, Gly64 in CesA, and Leu131 in both). Furthermore, the Nε2 atom of the catalytic His274 should be at hydrogen bonding distance to the oxygen of the substrate alcohol and to the catalytic Ser130 Oγ. In this configuration the substrate S-naproxenethylester binds with the alcohol moiety deep in the active site, and the naphtyl group at the hydrophobic entrance to the active site. Note, that the entrance to the active site is only 5.7 Å wide, measured between Cβ of Ala64 and Cζ of Phe166, and that it functions as a slit pocket. Using the same requirements, the model did not yield a productively bound R-naproxen ester substrate because of steric hindrance of the R-naproxen methyl group by Ala165, in agreement with the lack of activity of the enzyme towards this compound (Dröge et al., 2005).

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Figure 6. Stereo image of the modeling of the tetrahedral intermediate of (S)- (in cyan) and (R)-naproxen ester (in magenta) in the active site of NP. Amino acids Phe66 and A156, restricting access to the active site, are colored in yellow.

Applying the same criteria for productive binding, as used to model the naproxen ester in the active site of NP, the IPG moiety of the IPG esters most likely binds in the alcohol- binding pocket. The size of this pocket is mainly limited by Phe66 (Fig. 6), and could fit only methyl or ethyl substrates, in agreement with the enzyme's activity on naproxen methyl and ethyl esters (Dröge et al., 2005). However, the ability of the carboxylesterases to also hydrolyze the larger IPG esters (Dröge et al., 2001, Dröge et al., 2005) suggests that the active site alcohol-binding pocket is sufficiently flexible to accommodate the IPG-moiety. Only if Phe66, located at the interface between the two domains (Fig. 6A), adopts a different conformation, the alcohol-binding pocket would be able to accommodate the larger IPG alcohol moiety. Similar conformational changes have been observed in B. subtilis lipase (van Pouderoyen et al., 2001, Dröge et al., 2006) where, upon binding of an IPG-like inhibitor, the alcohol binding pocket expanded to accommodate the IPG moiety of the inhibitor. CesB is enantioselective towards IPG caprylate ester (Godinho et al., 2012), but NP is not. Indeed, our model shows that in CesB Tyr182 interacts with one of the IPG ring oxygen atoms, thus favoring the binding of the (S) enantiomer over the (R) enantiomer of IPG, where this hydrogen bond is not possible. In contrast, in NP the residue equivalent to Tyr182 is a Phe, which cannot make this hydrogen bonding interaction, resulting in a less selective binding. It is noteworthy that in the carboxylesterases NP, CesB and CesA no separate entry and exit routes (backdoor, side-door, or tunnel) for substrate and product are observed. The exit route found in other esterases (Cheeseman et al., 2004) is blocked by Phe162,

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His163, Pro164, Asp165 and Val166, all from the cap domain. Likewise, a possible water tunnel at the end of the alcohol-binding pocket is blocked by Phe66 (Fig. 3C, D). As a consequence, only one possible route to and from the active site is present in NP and CesB. However, in the reaction pathway the alcohol product is supposed to be the first to leave the active site (Scheme 1), but the exit route is blocked by the covalently bound acid part of the substrate. Similarly, the entrance route for the hydrolytic water molecule is obstructed. Yet, a lid-like movement of the cap domain (in particular of the flap made by helices α5 and α6) may be sufficient to allow the water molecule to enter the active site, and the products to be released. Indeed, the mainly hydrophobic interactions of the residues of helices α5 and α6 with the core domain could facilitate such a lid-like movement. Similar mechanisms of active site opening through cap domain helix relocation occur in family I.1 lipases (Nardini et al., 2000, Schrag et al., 1997) as well as in other α/β hydrolases (Otyepka et al., 2002, De Simone et al., 2004).

Comparison to other α/β-hydrolase fold esterases: similarity with meta-cleavage product hydrolases

A search of the Dali database (Holm et al., 2000) revealed that the NP and CesB structures are not very similar to those of other esterases, with the exception of the meta-cleavage product (MCP) hydrolases (Table S1). In general, the active site residues and oxyanion binding site amides are structurally conserved, but the alcohol-binding pockets of CesB and NP are smaller than those of esterases with known structure. In NP the acyl part of the substrate likely binds in the entrance to the active site as shown in Fig. 6. In contrast, in other α/β-hydrolase fold esterases the acyl part is usually bound in a deep hydrophobic pocket at the interface between the two domains (Liu et al., 2004, De Simone et al., 2004, Jansson et al., 2003). However, at that position in NP, Ala156 and Glu157, two residues located at the domain interface, prevent binding of substrates containing a large acyl moiety. Instead, the acyl moiety binds in the entrance to the active site. This is possible, because the helices in the cap domain have a different position, causing the active site entrance of NP and CesB to have a different location compared to other esterases. A secondary structure motif (SSM) search (Krissinel et al., 2004) of the PDB indicated significant structural similarity of the carboxylesterases NP and CesB with C-C bond breaking enzymes that belong to the MCP hydrolase family (Seah et al., 2007). These latter enzymes have 19-20% sequence identity to NP (Table S1) and overall root mean squared (r.m.s) deviations of less than 2.3 Å. This superfamily of cytosolic hydrolases also includes soluble non-heme peroxidases, epoxide hydrolases, fluoroacetate dehalogenases and haloalkane dehalogenases (Fischer et al., 2003). Several MCP hydrolase family members also show esterase activity (Alcaide et al., 2013). Intriguingly,

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compared to the dual specificity ester/MCP hydrolases, CesB and NP have an Ala156 insertion, which extends the loop after β6 and reduces the size of the active site, which probably contributes to their enantioselectivity. Carboxylesterases and MCP hydrolases share a common position of the cap domain. In some MCP hydrolases the α-helical lid (corresponding to residues 199-213 of CesB/NP) closes upon ligand binding (Lack et al., 2010) reducing the solvent accessibility of several hydrophobic residues. In NP and CesB residues 214-227, in the loop after helix α7, show the largest conformational differences compared to the MCP hydrolases (Fig. 3C). In fact, this is also the region where the structures of NP, CesA and CesB are most different. From these observations we conclude that the residues in this region may be flexible, which is also apparent from their elevated B-factor values. We suggest that this flexibility may be important to accommodate different substrates.

CONCLUSIONS

The structures of the B. subtilis carboxylesterases NP and CesB determined in the present study are very similar. Comparison of these structures to those of other α/β hydrolase fold esterases revealed that the main difference is the location of the entrance to active site, mainly because of a different positioning of the cap domain. In this respect NP and CesB resemble much more the MCP hydrolases, halogenases, dehalogenases and epoxide hydrolases than other esterases. Modeling of a substrate in the active site of NP indicates that the acyl chain is most likely bound at the entrance to the active site, unlike most other esterases, which have a well-defined pocket at the interface between the core and the cap domains (Jansson et al., 2003). Enantioselectivity of the carboxylesterases towards the IPG-substrates is mainly due to a few residues in the active site, Phe166 and Phe182 in NP vs Val166 and Tyr182 in CesB. Ala156, an insertion in the carboxylesterases, may be a determinant of enantioselectivity as well, since its side chain appears to interfere with the binding of certain enantiomers in the active site of NP as shown by the substrate modeling. In NP and CesB a lid domain movement for binding of the substrate and release of the product seems possible by movement of the flap made by helices α5 and α6 or/and of the loop after helix α7, where most of the amino acid and conformational differences between the three esterases occur. Such movements could also explain their activity on alcohol groups of the substrates that are too large to bind in the present conformation of the carboxylesterases. Altogether, these observations make the carboxylesterases unique members of the esterase family, with their own specific enantiospecificities, and indicate an evolutionary relationship to the soluble meta-cleavage compound hydrolases.

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ACKNOWLEDGEMENTS

We thank Johan Hekelaar for technical assistance with data collection of CesB crystals. We are grateful to the staff scientists at beam line BW7B at the EMBL-Hamburg outstation (DESY), Germany, for their support during data collection.

SUPPORTING MATERIAL

Figure S1. Structure–based alignment of CesB, NP and CesA. The structural alignment was made with SSM (http://www.ebi.ac.uk/msd-srv/ssm/). The secondary structure elements at the top and at the bottom of the sequence alignment are from the crystal structures of CesB and NP respectively. Identical residues have a red background color, similar residues have a red color. The figure was created with ESPript (Gouet et al., 1999).

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Figure S2. Secondary structure topology diagram representing Bacillus subtilis Thai I-8 NP and B. subtilis strain 168 CesB. Elements of the “canonical” core are indicated in light and dark grey. α-Helices (α) are represented by rectangles, 310-helices (η) by circles and β-strands (β) by arrows. The location of the active site is indicated by black dots and the corresponding amino acid residue. Differences between NP and CesB are indicated by italic labels NP and CesB

Table S1 Structural homology of NP to other enzymes, sorted by Z-score. Structural alignment is performed with DaliLite (Holm et al., 2000).

EC: EC number Z-score: statistical significance of a match in terms of Gaussian statistics. Rmsd: Rms deviation Id.: identity Nres: superposed Cα atoms MR: used in molecular replacement Ref.: reference

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PDB ID EC Z-score Rmsd Id. Nres MR Ref. NP carboxylesterase NP 4CCW 3.1.1.1 - - - 300 CesA carboxylesterase NA 2R11 3.1.1.1 45.1 1.2 98 300 - CesB (YbfK) 4CCY 3.1.1.1 45.1 1.2 64 296 CarC 2-hydroxy-6-oxo-6-(2(')- 1J1I 3.7.1.8 30.6 2.1 19 258 (Habe et aminophenyl)-hexa-2,4-dienoate al., 2003) hydrolase HsaD 2-hydroxy-6-oxo- 2VF2 3.7.1.8 29.9 2.3 25 282 (Lack et phenylhexa-2,4-dienoate al., 2008) hydrolase CumD 2-hydroxy-6-oxo-7- 1IUN 3.7.1.9 29.6 2.3 20 256 x (Fushinob methylocta-2,4-dienoate u et al., hydrolase 2002) ME7 CCSP0084 4I3F 29.5 2.2 21 253 (Alcaide et al., 2013) MhpC 2-hydroxy-6-ketonona- 1U2E 3.7.1.14 29.4 2.3 12 257 (Dunn et 2,4-diene-1,9-dioic acid 5,6- al., 2005) hydrolase BphD 2-hydroxyl-6-oxo-6- 1C4X 3.7.1.8 29.4 2.3 17 302 x (Horsman phenylhexa-2,4- dienoic acid et al., hydrolase 2006) RPA1163 fluoroacetate 3R3U 3.8.1.3 29.0 2.4 17 302 (Chan et dehalogenase al., 2011) PcaD 3-oxoadipate-enol- 2XUA 3.1.1.24 28.6 2.2 22 261 (Bains et lactonase al., 2011) E-2AMS (E)-2- 3KXP - 26.7 2.7 20 268 (McCulloc (acetamidomethylene) succinate h et al., hydrolase 2010) PFE Aryl esterase 1VA4 3.1.1.2 26.2 2.4 21 251 (Cheesema n et al., 2004) Ephy soluble epoxide hydrolase 1EHY 3.3.2.10 25.8 2.9 18 282 (Nardini et al., 1999b) EST pseudomonas putida 1ZOI 3.1 25.1 2.8 20 283 x (Elmi et al., 2005) MenH Stapylococcus aureus 2XMZ 4.2.99.20 24.5 2.7 17 238 (Dawson et al., 2011) RmdC Aclacinomycin 1Q0R -3.1 23.4 3.1 16 251 (Jansson et methylesterase al., 2003) BioH Pimelyl-[acyl-carrier 1M33 3.1..1.85 22.8 2.9 22 184 (Sanishvili protein] methyl ester esterase et al., 2003) EST30 Carboxylesterase 1TQH 3.1.1.1 23.2 2.2 18 176 (Liu et al., 2004)

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140

Chapter 7

Summary, General discussion & Outlook

......

The central dogma of molecular biology is that the sequence of the nucleic acids in (the coding sequence of) a gene specifies the structure and function of a protein, and that mutations in the nucleotide sequence may result in the evolution of protein variants with changed properties. An important and frequently observed feature of protein evolution is gene duplication, followed by divergence and/or domain fusion events. Following duplication, the original gene product retains its original function, but the duplicated gene product has no functional constraints and is allowed to evolve and develop new and different functions. Some homologous proteins may even diverge beyond the point of observable sequence similarity. In contrast, protein structures are much better conserved throughout evolution than the protein’s sequence. Hence, 3D structures allow for easier recognition of evolutionary relationships. Thus, to increase further understanding of the nature of evolutionary processes of proteins the combination of structural and sequence data will be helpful. This thesis discusses the crystal structures of six different enzymes. These six enzymes belong to different protein families and utilize different catalytic mechanisms. Understanding their 3D structures is essential for understanding the molecular determinants of their function and may shed a light on how they have evolved.

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MANNURONAN C-5-EPIMERASE

Alginate is synthesized by brown algae (Phaeophyceae, seaweed, kelp) and by Azotobacter and Pseudomonas species (Tondervik et al., 2013), and is a major constituent of mature biofilms produced by the latter bacteria. Alginate is industrially harvested all over the world. Commercially, alginates are interesting because of their gelling and viscosifying properties. Alginate is capable of absorbing 200-300 times its own weight in water, which makes it useful as an additive in products such as slimming aids, and for thickening drinks and cosmetics. It is also used for the water- and fireproofing of fabrics, for surface sizing in the paper industry and for textile printing. The most important application is probably in ice cream production, where alginates are used to prevent crystallization and shrinkage and to give a homogeneous product (Clementi, 1997). Alginate is a family of linear copolymers of (1→4)-linked β-D-mannuronic acid (M) and its C-5 epimer α-L-guluronic acid (G). The polymer is first produced as polymannuronic acid and the guluronic acid residues are then introduced at the polymer level as blocks by mannuronan C-5-epimerases. In Azotobacter vinelandii a family of 7 secreted and calcium-dependent, mannuronan C-5 epimerases (AlgE1-7) has been identified (Aachmann et al., 2006). These enzymes are highly homologous and consist of one or two catalytic A-modules and one to seven regulatory R-modules, of which at least one is needed for full activity (Ertesvåg et al., 1999). The mannuronan epimerases probably evolved through divergent evolution so that different epimerases introduce different G distribution patterns. The smallest of the AlgE proteins, AlgeE4, comprises one A- followed by one R-module and strictly forms alternating MG sequences (MG-blocks). The 2.1-Å resolution 3D structure of the catalytic A-module of the Azotobacter vinelandii mannuronan C-5-epimerase AlgE4 is described in chapter 2. AlgE4A folds into a right- handed parallel β-helix structure, originally found in pectate lyase C and subsequently in several other polysaccharide lyases and hydrolases. The AlgE4 β-helix is composed of four parallel β-sheets, comprising 12 complete turns, and has an amphipathic α-helix near the N terminus. The catalytic site is positioned in a positively charged cleft formed by loops extending from the surface encompassing Asp152, an amino acid previously shown to be important for the reaction. Site-directed mutagenesis further implicates Tyr149, His154, and Asp178 as being essential for activity. Tyr149 probably acts as the proton acceptor, whereas His154 is the proton donor in the epimerization reaction Interestingly, a striking agreement was noted in the spatial arrangement of these four amino acid residues in the center of the substrate binding clefts of alginate lyases from different polysaccharide lyase families (PL). AlgE4A displays the same spatial arrangement of Asp152, His154, Lys117, and Tyr149 in its active site as the equivalent Gln/Asn, His, Arg, and Tyr in the alginate lyases from family PL-7 (β-jelly roll), PL-5,

((α/α)6 barrel) and PL-18 (β-jelly roll) . This suggests that the catalytic mechanisms of

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Summary, General Discussion & Outlook ...... the four lyase families are the same. Recently, 3D structures of the periplasmic alginate epimerase form Pseudomonas aeruginosa (β-helix, PDB ID 4NK6 (Wolfram et al., 2014) and the exotype PL17 alginate lyase from Saccharophagus degredans ((α/α)6 barrel + β- sandwich, PDB ID 4NEI (Park et al., 2014)) were published. Both enzymes also show a similar conservation of the active site residues. This is a clear example of evolutionary convergent active sites, which evolved independently at least three times. The structure of the R-module of Alge4, a β-roll structure, was solved by NMR (Aachmann et al., 2006). The complete AlgE4 protein might form a single elongated structure with a long, positively charged patch for substrate binding (see the Figure below), in which the R-module appears as a natural extension of the A-module (Aachmann et al., 2006). However, the crystallization of the complete AlgE4 protein failed so far. Alternatively, electron microscopy or small angle X-ray scattering could validate this proposal. The research on alginates and mannuronan C-5-epimerases is continued at NTNU in Trondheim, Norway (Tondervik et al., 2013).

α-1,4 GLUCAN LYASE

α-1,4 Glucan lyase (GLase) was first purified and characterized by Shukun Yu (Yu et al., 1993) while purifying amylases from the red seaweed Gracilariopsis lemaneiformis. Subsequently, other GLases were isolated from the fungi Morchella costata, M. vulgaris and Peziza ostracoderma (Bojsen et al., 1999b). The enzyme degrades α-l,4-glucans to yield the keto-monosaccharide sugar 1,5-D- anhydrofructose (AF). This is the first reaction step in the anhydrofructose pathway, which is believed to be an alternative glycogen catabolic pathway of α-glucans, such as glycogen and starch (Yu et al., 2006). The physiological importance of the AF pathway in red algae is likely related to its involvement in stress responses associated with carbon starvation, removal of reactive oxygen species, and defense against infections by other organisms (Yu et al., 2006). Anhydrofructose is a promising compound for application as a modifier of gluten and gliadin proteins to enhance their solubility, and as a calorie-free

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sugar. In addition, AF has an antibrowning effect in green tea and is able to slow turbidity development in black currant wine (Yuan et al., 2005).

Glycogen

1,4-Glucan lyase

CH OH CH OH CH OH 2 AF reductase 2 AF reductase 2 O (NADPH) O (NADPH) O OH OH OH OH OH OH OH OH O 1,5-Anhydro-D-glucitol 1,5-Anhydro-D-fructose 1,5-Anhydro-D-mannitol

Cytochrome P450 Hexokinase Monooxygenase (ATP) H2O - CH2OPO3 CH2OH CH2OH O O O OH OH OH OH OH OH OH HO O 1,5-Anhydro-D-glucitol Ascopyrone M D-Mannose 6-phosphate

Dehydratase Ascopyrone D e hydratase tautomerase (isomerization) (isomerization)

H OH CH OH 2 O OH O OH CH2OH O CH2OH O OH O Ascopyrone P Microthecin 5-Epipentenomycin I

Scheme 1. The anhydrofructose pathway of glycogen catabolism (Yu, 2008).

The crystal structure of GLase, described in chapter 3, shows that similar to glycoside hydrolase family 31 (GH31) hydrolases, the enzyme contains a (β/α)8-barrel catalytic domain with B and B’ subdomains, an N-terminal domain N, and two C-terminal

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Summary, General Discussion & Outlook ...... domains, C and D. Domain N showed binding of a oligosaccharide, explaining its starch binding capacity. Furthermore, the 3D structures of GLase with inhibitors bound in the active site suggest that, as in GH31 hydrolases, the aspartic acid residues Asp553 and Asp665 are the catalytic nucleophile and acid, respectively. The active site of GLase superimposes well on those of other GH31 members. Our results show that the first step of the reaction, the formation of a covalent enzyme-substrate intermediate, is conserved in both lyases and hydrolases. Yet, the second step is different, since GLases catalyze a lyase reaction, and not a hydrolysis reaction as done by the GH31 glucan hydrolases. It appears that a subtle change, a Val to Glu substitution at position 556, promotes the lyase reaction. In GH31 hydrolases the strictly conserved Arg649 has a salt bridge interaction with the conserved Glu556, but in the GH31 lyases the equivalent residue is a Val or Thr. As a result, the arginine side chain in GLase can now take up a position to pick up the proton abstracted from the sugar’s C2 carbon atom by the correctly positioned carbonyl oxygen atom of Asp553 and relay it to the solvent. This suggests that residue 556 (being either a Glu or a Val) is a major determinant of the hydrolase vs lyase reaction specificity of GH31 enzymes. As a consequence, in GH31 lyases the Asp553 nucleophile has acquired a dual function, acting also as the base that abstracts the proton from the C2 atom of the -1 glucose residue. To support the putative dual role of the nucleophile we made a quadruple mutant, including the Glu323(556) to Val mutation, in the active site of the homologous GH31 α- glucosidase from Sulfolobus solfataricus. The mutant protein showed detectable lyase activity and a complete loss of glucosidase activity. Thus, indeed the E323V mutation in the glucosidase shifts the reaction from hydrolase toward lyase activity. This change in activity demonstrates the importance of position 323(556) for hydrolase/lyase activity in GH31, likely by modulating the conformation of the fully conserved arginine in the active site. Evolving the glucosidase into a genuine, full-activity lyase with a kcat similar to that of GLase will require further mutational studies. Likely, the GH31 enzymes appear to have evolved from a common ancestor comprising a domain N, a catalytic domain A, domain C and D. and have subsequently diverged. Domains B and B’, the most diverging domain among the GH31 members, make up the entrance to the active site and are important for substrate specificity. Whereas domain N contributes residues essential for oligosaccharide binding and maintenance of the active site architecture, the roles of the C-terminal domains are not clear (Ernst et al., 2006). It is surprising that with only single amino acid substitution a major change in reaction specificity can be achieved.

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PYRROLOQUINOLINE QUINONE DEPENDENT ALCOHOL DEHYDROGENASE

Pyrroloquinoline quinone dependent alcohol dehydrogenase (PQQ-ADH) from Pseudogluconobacter saccharoketogenes is a versatile dehydrogenase oxidizing various alcohols and sugars. The enzyme requires pyrroloquinoline quinone (PQQ) as a cofactor. As detailed in chapter 4 the structure of the PQQ-ADH consists of an eight-bladed β- propeller fold with the PQQ located in the center of the molecule near the 8-fold pseudosymmetry axis. The enzyme binds one PQQ and one Ca2+ ion like other PQQ- dependent methanol/ethanol dehydrogenases. However, three of the four ligands of the Ca2+ ion differ from those of related dehydrogenases and they come from different parts of the polypeptide chain. The only conserved amino acid residue is Asp333. This residue is proposed to be the catalytic base, which is supported by the bound substrate mimic polyethylene glycol. The active site is open to the solvent and easily accessible. This explains why PQQ-ADH can oxidize a broad range of substrates, extending for example from as small as methanol to as large as cyclodextrins. One of the closest homologues of PQQ-ADH is a type II Quinohemoprotein alcohol dehydrogenase QH-ADH (Oubrie et al., 2002), of which the structure was also determined in the Laboratory of Biophysical Chemistry. The largest difference between the two structures is that QH-ADH contains its own cytochrome c domain necessary for transfer of the electrons generated during the reaction. The relatively open active site of PQQ-ADH suggests that it can bind a cytochrome acceptor in a location that is similar to that of the cytochrome c domain of QH-ADH dehydrogenase. Another major difference of PQQ-ADH compared to type I Quinohemoprotein methanol dehydrogenases and type II enzymes is that PQQ-ADH lacks the vicinal disulfide bridge, implicated in electron transfer. The different calcium ligands, the absence of a cytochrome c domain, and the absence of the vicinal disulfide bond together create the open active site characteristic of this enzyme. Because of these major differences PQQ- ADH may be regarded as a phylogenetic outgroup in the tree of PQQ-dependent alcohol dehydrogenases. Probably the eight-bladed β-propeller PQQ dehydrogenases share a common ancestor. Over time, PQQ-ADH has accumulated differences compared to the other structurally characterized types, and has diverged into an enzyme with a very broad substrate specificity suitable for many applications.

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ENDO-XYLOGALACTURONAN HYDROLASE

Plant cell wall polysaccharides are the most abundant organic compounds found in nature. They make up 90% of the plant cell wall and can be divided into three groups: cellulose, hemicellulose, and pectin (de Vries et al., 2001). Pectins are heteropolysaccharides and consist of a backbone of α-1,4-linked D-galacturonic acid residues (polygalacturonan) decorated with various side chains or modifications. The “smooth” regions consist of polygalacturonan, of which the galacturonic acid residues may be acetylated at O-2 or O-3 or methylated at O-6. The “hairy” regions consist of xylogalacturonan (XGA), a D-xylose-substituted galacturonan backbone (de Vries et al., 2001). As XGA contributes to membrane fouling in the ultra-filtration process for fruit juice clarification, it is crucial that XGA is completely degraded during enzymatic treatment of the fruit pulp. Xylogalacturonan hydrolase (XghA) from Aspergillus tubingensis has been found to be an efficacious enzyme to degrade XGA (Zandleven et al., 2006, van der Vlugt- Bergmans et al., 2000). The enzyme belongs to glycoside hydrolase family 28 (GH28). In chapter 5 the crystal structure of endo-xylogalacturonan hydrolase (XghA) is described. XghA folds into a right-handed parallel β-helical structure comprising 10 complete turns with an active site that is located in an open groove/cleft on the surface of the protein. The high degree of structural conservation of the active site and catalytic apparatus, which is shared with polygalacturonases, indicates that cleavage of the substrate proceeds in essentially the same way as found for the other GH28 enzymes. However, the substrate-binding cleft is much wider than in other polygalacturonases, in agreement with its specificity for xylosylated substrates. The most extensive interactions appear to occur at subsite +2. Validation of the assigned subsites for XghA binding is a subject for future mutational experiments. XghA has probably evolved through gene duplication of an ancestral GH28 enzyme followed by divergent evolution. GH28 enzymes contain a β-helix structure observed in many carbohydrate-binding proteins and sugar hydrolases. The need for an extensive repertoire of enzymes in Aspergillus that can efficiently degrade pectin is illustrated by large amount of GH28 enzymes that have evolved. XghA plays a unique role in GH28 as it specifically degrades xylogalacturonan.

CARBOXYLESTERASES FROM BACILLUS SUBTILIS

Although the crystallization of naproxen esterase (NP) from Bacillus subtilis was already reported in 1993 (van der Laan et al., 1993), it took some 20 years to solve its structure (chapter 6). In 1999, diffraction data from a native NP crystal were collected by M.

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Nardini. However, no heavy atom derivatives for the enzyme could be found, and in the absence of close structural homologues, structure determination by molecular replacement was neither feasible. Only after an ensemble of different search models with less than 20% identity were used a partial structure was obtained consisting mainly of the central β-sheet structure of the α/β hydrolase fold. The structure determination of carboxylesterase CesB from Bacillus subtilis (chapter 6) was straightforward using NP as search model. NP and CesB show the typical α/β hydrolase structure that contains a catalytic triad comprising the active site. The structures consist of a core of 8 mostly parallel β-strands that are surrounded on both sides by α-helices, with an extra N-terminal helix stabilizing the cap subdomain. The catalytic triad is composed of Ser130, Glu245, and His274. The access/exit route for substrate and product to/from the active site and the solvent is not obvious from the structures. Flexibility of the cap domain might provide such an access/exit route. A Molecular Dynamics study may support this proposal. Surprisingly, the search models that were used for molecular replacement were not structures of esterases but of C-C bond breaking enzymes from the meta-cleavage product (MCP) hydrolase family. Despite their different activity, MCP hydrolases are most similar to the carboxylesterases. Recently, enzymes with dual esterase / MCP hydrolase activity were found (Alcaide et al., 2013), indicating that these enzymes show substrate promiscuity. It would be an interesting experiment to establish whether NP and CesB have promiscuous C-C bond cleaving activity.

CONCLUSION

In conclusion, the research described in this thesis has shed a light on the molecular details of the various evolutionary processes leading to divergence of reaction specificity of glucan lyase, divergence of substrate specificity of xylogalacturonan hydrolase and of PQQ-dependent alcohol dehydrogenase, divergence of enantioselectivity in carboxylesterases and to the convergent reaction specificity of mannuronan epimerases. How subtle changes in amino acid sequence lead to important changes in functionality is a continuous source of wonder that motivates crystallographers, and attests that evolution of a new function is still hard to predict.

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Nederlandse samenvatting

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Het centrale dogma van de moleculaire biologie is dat de nucleïnezuur (DNA) volgorde van een gen de structuur en functie van een eiwit specificeert, en dat mutaties in de DNA- volgorde kunnen leiden tot het ontstaan van eiwitvarianten met veranderde eigenschappen. Een belangrijke en vaak waargenomen gebeurtenis in de evolutie van eiwitten is gen duplicatie, gevolgd door divergentie en/of domein fusies. Na duplicatie behoudt één van beide gen producten zijn originele functie, maar het andere gen product heeft geen functionele beperking en kan vrij evolueren en nieuwe en andere functies ontwikkelen. Sommige eiwitten kunnen zover divergeren dat hun verwantschap niet meer in hun aminozuur volgorde waarneembaar is. Daarentegen, blijven de drie- dimensionale structuren van de eiwitten veel beter bewaard dan de aminozuur volgordes tijdens de evolutie. Daarom kunnen evolutionaire verwantschappen eenvoudiger in 3D-structuren worden herkend. Dus, om verdere kennis van de evolutionaire ontwikkelingen van eiwitten te verkrijgen zal de combinatie van structurele en volgorde gegevens nuttig zijn. Inzicht in hoe enzymen zich ontwikkeld hebben als biologische katalysatoren om de grote verscheidenheid aan reacties uit te voeren wordt geïllustreerd met zeven (klassieke) voorbeelden in hoofdstuk 1. De driedimensionale structuren van deze eiwitten en hun evolutionaire relaties met andere eiwitten en enzymen worden besproken. Dit proefschrift beschrijft de kristalstructuren van zes verschillende enzymen. Deze zes enzymen behoren tot verschillende eiwitfamilies en maken gebruik van verschillende katalytische mechanismes. Begrip van hun 3D-structuren is essentieel voor het begrijpen van de moleculaire basis van hun functioneren en kan hun evolutie aanzienlijk verhelderen. De 2.1 Å resolutie 3D structuur van de katalytische A-module van Azotobacter vinelandii mannuronan C-5-epimerase AlgE4 wordt beschreven in hoofdstuk 2. AlgE4A vouwt als een rechtshandige parallelle -helix structuur, oorspronkelijk gevonden in pectate lyase C en vervolgens in verschillende andere polysaccharide lyasen en hydrolasen. De AlgE4 β-helix heeft 12 complete β-strand windingen, die samen vier parallelle β-sheets vormen; de β-helix heeft een amfipatische α-helix nabij het N-terminale uiteinde. Het katalytisch centrum is gesitueerd in een positief geladen groef die gevormd wordt door peptide ketens die uit het β-helix oppervlak steken. Hier bevindt zich ook Asp152, een aminozuur waarvan eerder is aangetoond dat het belangrijk is voor de reactie. Verder geeft plaatsgerichte mutagenese (site-directed mutagenesis) aan dat Tyr149, His154 en Asp178 essentieel zijn voor de activiteit. Waarschijnlijk werkt Tyr149 als de proton acceptor, terwijl His154 de proton donor is in de epimerizatie reactie. Een opvallende overeenkomst werd opgemerkt in de ruimtelijke rangschikking van deze vier aminozuren in het midden van de substraat bindingsgroeven van alginaat lyasen uit verschillende polysaccharide lyase families (PL). AlgE4A laat dezelfde ruimtelijke ordening zien van Asp152, His154, Lys117, en Tyr149 in zijn actief centrum als de equivalente

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Gln/Asn, His, Arg, en Tyr in de alginaat lyasen van familie PL-7 (β-jelly rol), PL-5, ((α/α)6 barrel) en PL-18 (β-jelly rol). Dit suggereert dat de katalytische mechanismen van de vier lyase families hetzelfde zijn. Dit is een duidelijk voorbeeld van evolutionair convergente actieve centra die ten minste driemaal onafhankelijk van elkaar zijn ontstaan. De kristalstructuur van α-1,4 Glucan lyase (GLase) van het rode zeewier Gracilariopsis lemaneiformis, beschreven in hoofdstuk 3, laat zien dat het enzym bestaat uit een

(β/α)8-barrel katalytisch domein met B en B’ subdomeinen, een N-terminaal domein, en twee C-terminale domeinen, C en D. Domein N bleek een oligosaccharide te binden, hetgeen het zetmeel bindend vermogen van het enzym verklaart. De 3D structuur van GLase is zeer vergelijkbaar met die van enzymen behorend bij de GH31 glycoside hydrolase familie, maar GLase is geen hydrolase maar een lyase. 3D structuren van GLase met gebonden remmers laten zien dat, net als in de GH31 hydrolasen, de aspartaten D553 en D665, respectievelijk het katalytische nucleofiel en zuur zijn. Bovendien superponeert het actief centrum van GLase goed op dat van andere GH31 leden. Onze resultaten laten zien dat de eerste stap in de reactie, de vorming van een covalent enzym-substraat intermediair geconserveerd is in zowel de lyasen en hydrolasen. Echter, de tweede stap is verschillend, omdat GLasen een lyase reactie katalyseren en geen hydrolase reactie zoals gedaan wordt door GH31 glucan hydrolasen. Het blijkt dat een subtiele verandering, een Val naar Glu vervanging op positie 556, de lyase reactie bevordert. In GH31 hydrolasen heeft de strikt geconserveerde R649 een zoutbrug interactie met de geconserveerde E556, maar in GH31 lyasen is het equivalente aminozuur een Val of Thr. Als gevolg daarvan kan de R649 zijketen in GLase een andere positie aannemen die geschikt is om het proton op te pikken dat de D553 carboxylaat van het C2 koolstofatoom van het substraat heeft gehaald, en dit door te geven aan het oplosmiddel. Dit suggereert dat aminozuur 556 (hetzij een Glu of een Val) een belangrijke factor is voor de hydrolase vs. de lyase reactie specificiteit van GH31 enzymen. Bovendien heeft de D553 carboxylaat een dubbele functie gekregen, want behalve dat het fungeert als nucleofiel is het ook de base die het proton van het C2- atoom van het substraat haalt. Om de mogelijke dubbelrol van de nucleofiel aan te tonen hebben we een viervoudige mutant gemaakt, met daarin de Glu323 (556) naar Val mutatie, in het actieve centrum van de homologe GH31 α-glucosidase uit Sulfolobus solfataricus. Het gemuteerde eiwit vertoonde detecteerbare lyase activiteit en een volledig verlies van glucosidase activiteit. Dus verschuift de E323V mutatie inderdaad de reactie van hydrolase naar lyase activiteit. Ons werk heeft dus laten zien hoe een subtiele aminozuur verandering een volledig nieuwe activiteit kan den ontstaan. Pyrroloquinoline quinone afhankelijk alcohol dehydrogenase (PQQ-ADH) van Pseudogluconobacter saccharoketogenes is een veelzijdige dehydrogenase die diverse

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alcoholen en suikers oxideert. Het enzym heeft pyrroloquinoline quinone (PQQ) nodig als cofactor. Zoals beschreven in hoofdstuk 4 bestaat de structuur van PQQ-ADH uit een acht-bladige β-propeller vouwing met PQQ gesitueerd in het centrum van het molecuul nabij de as van 8-voudige pseudosymmetrie. Het enzym bindt een PQQ en een Ca2+ ion net zoals andere PQQ-afhankelijke methanol/ethanol dehydrogenasen. Echter, drie van de vier liganden van het Ca2+ ion verschillen van die van verwante dehydrogenasen en ze komen van verschillende delen van de polypeptide keten. Het enige geconserveerde aminozuur is Asp333. Dit aminozuur is waarschijnlijk de katalytische base, wat ondersteund wordt door een gebonden substraat analoog (polyethyleenglycol). Het actief centrum is gemakkelijk toegankelijk en wijd open. Dit verklaart waarom PQQ-ADH een groot aantal substraten, van bijvoorbeeld zo klein als methanol tot zo groot als cyclodextrines kan oxideren. Het grote verschil met de verwante methanol dehydrogenasen is dat PQQ-ADH de zwavel brug mist die betrokken is bij elektronen overdracht. Een ander groot verschil is de afwezigheid van het cytochroom c domein dat noodzakelijk is voor de elektronen overdracht. Er is echter ruimte genoeg om een los cytochroom eiwit te binden. Al deze verschillen maken PQQ-ADH een bijzonder intrigerend evolutionair buitenbeentje. In hoofdstuk 5 wordt de kristalstructuur van endo-xylogalacturonan hydrolase (XghA) beschreven. XghA hoort bij glycoside hydrolase familie 28 (GH28) en speelt een unieke rol in deze familie aangezien het specifiek xylogalacturonan afbreekt. XghA vouwt als een rechtshandige parallelle β-helix structuur bestaand uit 10 complete windingen met een actief centrum dat is gelegen in een open groef/kloof op het oppervlak van het eiwit. De hoge mate van structurele conservering van het actief centrum en de katalytische inrichting, die wordt gedeeld met polygalacturonases, geeft aan dat de substraat omzetting in principe op dezelfde wijze verloopt als bij andere GH28 enzymen. Echter, de substraat bindende groef is veel breder dan in andere polygalacturonasen, in overeenstemming met zijn specificiteit voor gexylosyleerde substraten. De meest uitgebreide interacties lijken voor te komen bij subsite +2. Validering van de gedefinieerde subsites in XghA binding is een onderwerp voor toekomstige mutatie experimenten. In hoofdstuk 6 worden de kristalstructuren van de carboxylesterases naproxen esterase (NP) en CesB beschreven. NP en CesB vertonen de typische α/β hydrolase opvouwing met drie katalytische aminozuren, Ser130, Glu245, en His274 als actief centrum. De structuren bestaan uit een centraal domein van 8 grotendeels parallelle β-strands die aan beide zijden omgeven zijn door α-helices, met een extra N-terminale helix die het deksel subdomein stabiliseert. De toegangs/uitgangs route voor substraat en product naar/van het actief centrum en het oplosmiddel is niet duidelijk te vinden in de structuren. Flexibiliteit van het deksel domein zou in die toegangs/uitgangs route kunnen voorzien. Ondanks hun verschillende activiteiten zijn de carboxylesterases

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Nederlandse samenvatting ...... structureel het meest homoloog aan enzymen die C-C bindingen breken, de "meta- cleavage product” (MCP) hydrolasen. Samengevat, heeft het in dit proefschrift beschreven onderzoek meer duidelijkheid gegeven over de moleculaire details van de verschillende evolutionaire processen die leiden tot divergentie van de reactie specificiteit van enzymen (glucan lyase), divergentie van de substraat specificiteit (xylogalacturonan hydrolase, PQQ-afhankelijke alcohol dehydrogenase), divergentie van enantioselectiviteit (carboxylesterases), en de convergente reactie specificiteit van mannuronan epimerasen. Hoe subtiele veranderingen in de aminozuur volgorde leiden tot belangrijke wijzigingen in functionaliteit is een voortdurende bron van verwondering (en motivatie!) voor kristallografen, en geeft aan dat de evolutie van een nieuwe functie nog steeds moeilijk te voorspellen is.

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De auteur van dit proefschrift werd geboren te Appingedam op 12 juli 1964. Aansluitend op het behalen van haar eindexamen VWO aan het Ommelander College in Appingedam, is zij in 1982 begonnen met haar studie Hoger Laboratorium Onderwijs (Chemie) aan de Dagschool voor Laboratoriumpersoneel in Groningen. Zij liep een 1-jarige stage bij de Keuringsdienst van Waren in Groningen alwaar zij voornamelijk onderzoek deed naar het bepalen van biologische amines in vis met behulp van HPLC technieken. Na het behalen van het diploma HLO is zij aan de slag gegaan als analist in het O-lab van Akzo- Nobel te Delfzijl. In oktober 1986 is zij gaan werken als analiste in de vakgroep Chemische Fysica – Biomoleculen geleid door hoogleraar W. Hol. Zij werd aangesteld op het Protein Engineering project onder begeleiding van B.W. Dijkstra. Op 1 juli 1993 werd zij in vaste dienst benoemd. Tijdens 28 jaren heeft zij aan een groot aantal projecten gewerkt, voornamelijk eiwitzuivering en kristallisatie, en in de laatste 15 jaren ook eiwit structuur opheldering. Enige projecten worden in dit proefschrift beschreven. Sinds 1 augustus jl. werkt zij als eiwitkristallografe in de Biotransformation and Biocatalysis groep van Prof. D.B. Janssen en Prof. M.W. Fraaije.

1. Rozeboom, H. J., Kingma, J., Janssen, D. B., and Dijkstra, B. W. (1988) Crystallization of haloalkane dehalogenase from xanthobacter-autotrophicus GJ10. J. Mol. Biol. 200, 611-612. 2. Vanderklei, I. J., Lawson, C. L., Rozeboom, H., Dijkstra, B. W., Veenhuis, M., Harder, W., and Hol, W. G. J. (1989) Use of electron-microscopy in the examination of lattice-defects in crystals of alcohol oxidase. FEBS Lett. 244, 213-216. 3. Rozeboom, H. J., Budiani, A., Beintema, J. J., and Dijkstra, B. W. (1990a) Crystallization of hevamine, an enzyme with lysozyme chitinase activity from hevea-brasiliensis latex. J. Mol. Biol. 212, 441-443. 4. Rozeboom, H. J., Dijkstra, B. W., Engel, H., and Keck, W. (1990b) Crystallization of the soluble lytic transglycosylase from escherichia-coli K12. J. Mol. Biol. 212, 557-559. 5. Franken, S. M., Rozeboom, H. J., Kalk, K. H., and Dijkstra, B. W. (1991) Crystal-structure of haloalkane dehalogenase - an enzyme to detoxify halogenated alkanes. EMBO J. 10, 1297-1302. 6. Verschueren, K. H. G., Seljee, F., Rozeboom, H. J., Kalk, K. H., and Dijkstra, B. W. (1993a) Crystallographic analysis of the catalytic mechanism of haloalkane dehalogenase. Nature. 363, 693-698. 7. Verschueren, K. H. G., Kingma, J., Rozeboom, H. J., Kalk, K. H., Janssen, D. B., and Dijkstra, B. W. (1993b) Crystallographic and fluorescence studies of the interaction of haloalkane dehalogenase with halide-ions - studies with halide compounds reveal a halide binding-site in the active-site. Biochemistry (N. Y.). 32, 9031-9037. 8. Verschueren, K. H. G., Franken, S. M., Rozeboom, H. J., Kalk, K. H., and Dijkstra, B. W. (1993c) Noncovalent binding of the heavy-atom compound [Au(CN)2]- at the halide binding-site of haloalkane dehalogenase from xanthobacter-autotrophicus GJ10. FEBS Lett. 323, 267-270.

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9. Verschueren, K. H. G., Franken, S. M., Rozeboom, H. J., Kalk, K. H., and Dijkstra, B. W. (1993d) Refined X- ray structures of haloalkane dehalogenase at pH 6.2 and pH 8.2 and implications for the reaction- mechanism. J. Mol. Biol. 232, 856-872. 10. Terwisscha van Scheltinga, A. C., Rozeboom, H. J., Kalk, K. H., and Dijkstra, B. W. (1993) X-Ray Studies on Hevamin, an Enzyme from Hevea Brasiliens with Chitinase and Lysozyme Activity, in Proceeding of the International Sympsium on Chitin Enzymology pp 205-210, . 11. Thunnissen, A. M. W. H., Dijkstra, A. J., Kalk, K. H., Rozeboom, H. J., Engel, H., Keck, W., and Dijkstra, B. W. (1994) Doughnut-shaped structure of a bacterial muramidase revealed by X-ray crystallography. Nature. 367, 750-754. 12. Lawson, C. L., Vanmontfort, R., Strokopytov, B., Rozeboom, H. J., Kalk, K. H., Devries, G. E., Penninga, D., Dijkhuizen, L., and Dijkstra, B. W. (1994) Nucleotide-sequence and X-ray structure of cyclodextrin glycosyltransferase from bacillus-circulans strain-251 in a maltose-dependent crystal form. J. Mol. Biol. 236, 590-600. 13. Dijkhuizen, L., Penninga, D., Rozeboom, H. J., Strokopytov, B., and Dijkstra, B. W. (1995) Protein Engineering of Cyclodextrin Glycosyltransferase from Bacillus Circulans Strain 251, in Proc. 3rd Int. Symp. Perspectives on Protein Engineering pp 96-99 . 14. Ridder, I. S., Rozeboom, H. J., Kingma, J., Janssen, D. B., and Dijkstra, B. W. (1995) Crystallization and preliminary-X-ray analysis of L-2-haloacid dehalogenase from xanthobacter-autotrophicus Gj10. Protein Science. 4, 2619-2620. 15. Knegtel, R. M. K., Strokopytov, B., Penninga, D., Faber, O. G., Rozeboom, H. J., Kalk, K. H., Dijkhuizen, L., and Dijkstra, B. W. (1995) Crystallographic studies of the interaction of cyclodextrin glycosyltransferase from bacillus-circulans strain-251 with natural substrates and products. J. Biol. Chem. 270, 29256-29264. 16. Dijkhuizen, L., Penninga, D., Rozeboom, H. J., Strokopytov, B., and Dijkstra, B. W. (1995) Protein Engineering of Cyclodextrin Glycosyltransferase from Bacillus Circulans Strain 251 in Progress in Biotechnology 10, Carbohydrate Bioengineering pp 165-174, . 17. Penninga, D., Strokopytov, B., Rozeboom, H. J., Lawson, C. L., Dijkstra, B. W., Bergsma, J., and Dijkhuizen, L. (1995) Site-directed mutations in tyrosine-195 of cyclodextrin glycosyltransferase from bacillus-circulans strain-251 affect activity and product specificity. Biochemistry (N. Y.). 34, 3368-3376. 18. Thunnissen, A. M. W. H., Rozeboom, H. J., Kalk, K. H., and Dijkstra, B. W. (1995) Structure of the 70-kda soluble lytic transglycosylase complexed with bulgecin-a - implications for the enzymatic mechanism. Biochemistry (N. Y.). 34, 12729-12737. 19. Strokopytov, B., Penninga, D., Rozeboom, H. J., Kalk, K. H., Dijkhuizen, L., and Dijkstra, B. W. (1995) X- ray structure of cyclodextrin glycosyltransferase complexed with acarbose - implications for the catalytic mechanism of glycosidases. Biochemistry (N. Y.). 34, 2234-2240. 20. Knegtel, R. M. A., Wind, R. D., Rozeboom, H. J., Kalk, K. H., Buitelaar, R. M., Dijkhuizen, L., and Dijkstra, B. W. (1996) Crystal structure at 2.3 angstrom resolution and revised nucleotide sequence of the thermostable cyclodextrin glycosyltransferase from thermoanaerobacterium thermosulfurigenes EM1. J. Mol. Biol. 256, 611-622. 21. Krengel, U., Rozeboom, H. J., Kalk, K. H., and Dijkstra, B. W. (1996) Crystallization and preliminary crystallographic analysis of endo-1,4-beta-xylanase I from aspergillus niger. Acta Crystallogr. D. 52, 571- 576. 22. Penninga, D., vanderVeen, B. A., Knegtel, R. M. A., vanHijum, S. A. F. T., Rozeboom, H. J., Kalk, K. H., Dijkstra, B. W., and Dijkhuizen, L. (1996) The raw starch binding domain of cyclodextrin glycosyltransferase from bacillus circulans strain 251. J. Biol. Chem. 271, 32777-32784. 23. Uitdehaag, J. C. M., Penninga, D., Alebeek, G., vanderVeen, B., Rozeboom, H., Kalk, K. H., Dijkhuizen, L., and Dijkstra, B. W. (1996) Residues in cyclodextrin glycosyl transferase that determine product specificity. Prog. Biophys. Mol. Biol. 65, PA130-PA130. 24. Strokopytov, B., Knegtel, R. M. A., Penninga, D., Rozeboom, H. J., Kalk, K. H., Dijkhuizen, L., and Dijkstra, B. W. (1996) Structure of cyclodextrin glycosyltransferase complexed with a maltononaose inhibitor at 2.6 angstrom resolution. implications for product specificity. Biochemistry (N. Y.). 35, 4241- 4249.

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25. Ridder, I. S., Rozeboom, H. J., Kalk, K. H., Janssen, D. B., and Dijkstra, B. W. (1997) Three-dimensional structure of L-2-haloacid dehalogenase from xanthobacter autotrophicus GJ10 complexed with the substrate-analogue formate. J. Biol. Chem. 272, 33015-33022. 26. Krooshof, G. H., Ridder, I. S., Tepper, A. W. J. W., Vos, G. J., Rozeboom, H. J., Kalk, K. H., Dijkstra, B. W., and Janssen, D. B. (1998) Kinetic analysis and X-ray structure of haloalkane dehalogenase with a modified halide-binding site. Biochemistry (N. Y.). 37, 15013-15023. 27. Oubrie, A., Rozeboom, H. J., Kalk, K. H., Duine, J. A., and Dijkstra, B. W. (1999a) The 1.7 angstrom crystal structure of the apo form of the soluble quinoprotein glucose dehydrogenase from acinetobacter calcoaceticus reveals a novel internal conserved sequence repeat. J. Mol. Biol. 289, 319-333. 28. Oubrie, A., Rozeboom, H. J., and Dijkstra, B. W. (1999b) Active-site structure of the soluble quinoprotein glucose dehydrogenase complexed with methylhydrazine: A covalent cofactor-inhibitor complex. Proc. Natl. Acad. Sci. U. S. A. 96, 11787-11791. 29. Ridder, I. S., Rozeboom, H. J., Kalk, K. H., and Dijkstra, B. W. (1999) Crystal structures of intermediates in the dehalogenation of haloalkanoates by L-2-haloacid dehalogenase. J. Biol. Chem. 274, 30672-30678. 30. Pikkemaat, M. G., Ridder, I. S., Rozeboom, H. J., Kalk, K. H., Dijkstra, B. W., and Janssen, D. B. (1999) Crystallographic and kinetic evidence of a collision complex formed during halide import in haloalkane dehalogenase. Biochemistry (N. Y.). 38, 12052-12061. 31. Ridder, I. S., Rozeboom, H. J., and Dijkstra, B. W. (1999) Haloalkane dehalogenase from xanthobacter autotrophicus GJ10 refined at 1.15 angstrom resolution. Acta Crystallogr. D. 55, 1273-1290. 32. Oubrie, A., Rozeboom, H. J., Kalk, K. H., Olsthoorn, A. J. J., Duine, J. A., and Dijkstra, B. W. (1999) Structure and mechanism of soluble quinoprotein glucose dehydrogenase. EMBO J. 18, 5187-5194. 33. Nardini, M., Ridder, I. S., Rozeboom, H. J., Kalk, K. H., Rink, R., Janssen, D. B., and Dijkstra, B. W. (1999) The X-ray structure of epoxide hydrolase from agrobacterium radiobacter AD1 - an enzyme to detoxify harmful epoxides. J. Biol. Chem. 274, 14579-14586. 34. Nardini, M., Ridder, I. S., Rozeboom, H. J., Kalk, K. H., Rink, R., Janssen, D. B., and Dijkstra, B. W. (2000) X-ray structure of epoxide hydrolase from agrobacterium radiobacter AD1: An enzyme to detoxify harmful epoxides. Ital. J. Biochem. 49, 15-15. 35. Oubrie, A., Huizinga, E. G., Rozeboom, H. J., Kalk, K. H., de Jong, G. A. H., Duine, J. A., and Dijkstra, B. W. (2001) Crystallization of quinohaemoprotein alcohol dehydrogenase from comamonas testosteroni: Crystals with unique optical properties. Acta Crystallogr. D. 57, 1732-1734. 36. Steiner, R. A., Rozeboom, H. J., de Vries, A., Kalk, K. H., Murshudov, G. N., Wilson, K. S., and Dijkstra, B. W. (2001) X-ray structure of bovine pancreatic phospholipase A(2) at atomic resolution. Acta Crystallogr. D. 57, 516-526. 37. Oubrie, A., Rozeboom, H. J., Kalk, K. H., Huizinga, E. G., and Dijkstra, B. W. (2002) Crystal structure of quinohemoprotein alcohol dehydrogenase from comamonas testosteroni - structural basis for substrate oxidation and electron transfer. J. Biol. Chem. 277, 3727-3732. 38. Fusetti, F., Schroter, K. H., Steiner, R. A., van Noort, P. I., Pijning, T., Rozeboom, H. J., Kalk, K. H., Egmond, M. R., and Dijkstra, B. W. (2002) Crystal structure of the copper-containing quercetin 2,3- dioxygenase from aspergillus japonicus. Structure. 10, 259-268. 39. de Jong, R. M., Rozeboom, H. J., Kalk, K. H., Tang, L. X., Janssen, D. B., and Dijkstra, B. W. (2002) Crystallization and preliminary X-ray analysis of an enantioselective halohydrin dehalogenase from agrobacterium radiobacter AD1. Acta Crystallogr. D. 58, 176-178. 40. Bokma, E., Rozeboom, H. J., Sibbald, M., Dijkstra, B. W., and Beintema, J. J. (2002) Expression and characterization of active site mutants of hevamine, a chitinase from the rubber tree hevea brasiliensis. Eur. J. Biochem. 269, 893-901. 41. Leemhuis, H., Uitdehaag, J. C. M., Rozeboom, H. J., Dijkstra, B. W., and Dijkhuizen, L. (2002) The remote substrate binding subsite-6 in cyclodextrin-glycosyltransferase controls the transferase activity of the enzyme via an induced-fit mechanism. J. Biol. Chem. 277, 1113-1119. 42. Fusetti, F., von Moeller, H., Houston, D., Rozeboom, H. J., Dijkstra, B. W., Boot, R. G., Aerts, J. M. F. G., and van Aalten, D. M. F. (2002) Structure of human chitotriosidase - implications for specific inhibitor design and function of mammalian chitinase-like lectins. J. Biol. Chem. 277, 25537-25544.

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43. Leemhuis, H., Rozeboom, H. J., Wilbrink, M., Euverink, G. J. W., Dijkstra, B. W., and Dijkhuizen, L. (2003a) Conversion of cyclodextrin glycosyltransferase into a starch hydrolase by directed evolution: The role of alanine 230 in acceptor subsite+1. Biochemistry (N. Y.). 42, 7518-7526. 44. Leemhuis, H., Rozeboom, H. J., Dijkstra, B. W., and Dijkhuizen, L. (2003b) The fully conserved asp residue in conserved sequence region I of the alpha-amylase family is crucial for the catalytic site architecture and activity. FEBS Lett. 541, 47-51. 45. de Jong, R. M., Tiesinga, J. J. W., Rozeboom, H. J., Kalk, K. H., Tang, L., Janssen, D. B., and Dijkstra, B. W. (2003) 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. EMBO J. 22, 4933-4944. 46. Leemhuis, H., Rozeboom, H. J., Dijkstra, B. W., and Dijkhuizen, L. (2004) Improved thermostability of bacillus circulans cyclodextrin glycosyltransferase by the introduction of a salt bridge. Proteins. 54, 128- 134. 47. Kelly, R. M., Leemhuis, H., Rozeboom, H. J., van Oosterwijk, N., Dijkstra, B. W., and Dijkhuizen, L. (2008) Elimination of competing hydrolysis and coupling side reactions of a cyclodextrin glucanotransferase by directed evolution. Biochem. J. 413, 517-525. 48. Rozeboom, H. J., Bjerkan, T. M., Kalk, K. H., Ertesvåg, H., Holtan, S., Aachmann, F. L., Valla, S., and Dijkstra, B. W. (2008) Structural and mutational characterization of the catalytic A-module of the mannuronan C-5-epimerase AlgE4 from azotobacter vinelandii. J. Biol. Chem. 283, 23819-23828. 49. Patel, A. K., van Oosterwijk, N., Singh, V. K., Rozeboom, H. J., Kalk, K. H., Siezen, R. J., Jagannadham, M. V., and Dijkstra, B. W. (2009) Crystallization and preliminary X-ray analysis of carnein, a serine protease from ipomoea carnea. Acta Crystallogr. F. 65, 383-385. 50. Wasiel, A. A., Rozeboom, H. J., Hauke, D., Baas, B., Zandvoort, E., Quax, W. J., Thunnissen, A. W. H., and Poelarends, G. J. (2010) Structural and functional characterization of a macrophage migration inhibitory factor homologue from the marine cyanobacterium prochlorococcus marinus. Biochemistry (N. Y.). 49, 7572-7581. 51. Ismaya, W. T., Rozeboom, H. J., Weijn, A., Mes, J. J., Fusetti, F., Wichers, H. J., and Dijkstra, B. W. (2011a) Crystal structure of agaricus bisporus mushroom tyrosinase: Identity of the tetramer subunits and interaction with tropolone. Biochemistry (N. Y.). 50, 5477-5486. 52. Ismaya, W. T., Rozeboom, H. J., Schurink, M., Boeriu, C. G., Wichers, H., and Dijkstra, B. W. (2011b) Crystallization and preliminary X-ray crystallographic analysis of tyrosinase from the mushroom agaricus bisporus. Acta Crystallogr. F. 67, 575-578. 53. Raj, H., Szymanski, W., de Villiers, J., Rozeboom, H. J., Veetil, V. P., Reis, C. R., de Villiers, M., Dekker, F. J., de Wildeman, S., Quax, W. J., Thunnissen, A. W. H., Feringa, B. L., Janssen, D. B., and Poelarends, G. J. (2012) Engineering methylaspartate ammonia lyase for the asymmetric synthesis of unnatural amino acids. Nature Chemistry. 4, 478-484. 54. Godinho, L. F., Reis, C. R., Rozeboom, H. J., Dekker, F. J., Dijkstra, B. W., Poelarends, G. J., and Quax, W. J. (2012) Enhancement of the enantioselectivity of carboxylesterase A by structure-based mutagenesis. J. Biotechnol. 158, 36-43. 55. Rozeboom, H. J., Yu, S., Madrid, S., Kalk, K. H., Zhang, R., and Dijkstra, B. W. (2013a) Crystal structure of alpha-1,4-glucan lyase, a unique glycoside hydrolase family member with a novel catalytic mechanism. J. Biol. Chem. 288, 26764-26774. 56. Rozeboom, H. J., Beldman, G., Schols, H. A., and Dijkstra, B. W. (2013b) Crystal structure of endo- xylogalacturonan hydrolase from aspergillus tubingensis. Febs Journal. 280, 6061-6069. 57. Xu, J., Chen, B., Callis, P., Rozeboom, H., Broos, J., and Knutson, J. (2013) Femtosecond fluorescence dynamics of tryptophan and 5-fluorotryptophan in monellin: Slow water relaxation unmasked. Biophys. J. 104, 681A-681A. 58. Rozeboom, H. J., Godinho, L. F., Nardini, M., Quax, W. J., and Dijkstra, B. W. (2014) Crystal structures of two bacillus carboxylesterases with different enantioselectivities. Biochim. Biophys. Acta. 1844, 567-575.

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Dankwoord

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Het is zover, de stof van mijn bureaustoel is doorgesleten, maar het boekje is af.

Hierbij wil ik iedereen bedanken die op enigerwijze een bijdrage heeft geleverd aan het tot stand komen van dit proefschrift. Een aantal mensen wil ik met naam noemen.

Als eerste mijn promotor Bauke Dijkstra. Bauke, hartelijk bedankt voor het vertrouwen in me. Vooral bedankt voor het corrigeren van de manuscripten. Het is bijzonder hoe jij de wetenschappelijke resultaten in heldere beknopte zinnen kunt vertalen.

Verder bedank ik de leescommissie Prof. T Sixma, Prof. G.J. Poelarends en Prof D.B. Janssen voor het beoordelen van mijn proefschrift.

I would like to thank Tonje Bjerkan, Helga Ertesvåg, Synnøve Holtan, Finn Aachmann and Svein Valla from Trondheim for the collaboration on the mannuronan epimerase. Tonje, I will never forget the week that you stayed in Groningen. You were here at a very special moment for me.

Of course, I would like to express my gratitude to Shukun Yu for providing me with two very interesting enzymes, Glucan lyase became my pet project too. I would also like to thank Susan Madrid for work on the MalA mutants and Ran Zhang for providing the fluoro sugars. Furthermore, I want to thank Rene Mikkelsen, Igor Nikolaev and Harm Mulder for their work on the quinone dependent alcohol dehydrogenase.

Verder wil ik Gerrit Beldman en Henk Schols bedanken voor de samenwerking met het endo-xylogalacturonan hydrolase project.

Luis Godinho and Wim Quax, thank you for the collaboration on the CesB enzyme. I would like to thank Marco Nardini for support with the naproxen esterase.

Dan wil ik al mijn(ex-)collega’s van de eiwitkristallografie en elektronen microscopie bedanken voor de fijne samenwerking en prettige sfeer op het lab. Jullie waren vast wel eens zat van het eeuwige Skyradio op het lab. In 28 jaar zijn er zoveel collega’s geweest dat ik ze niet allemaal persoonlijk kan bedanken. De collega’s van dit moment wil ik bedanken vooral voor de steun bij de laatste loodjes; Andy, Anke, Carol, Eswar, Harsh, Hilda, Ilona, Jaap, Jinfeng, Marco en Niels.

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Kor Kalk en Johan wil ik bedanken voor de assistentie bij het meten van de diffractie data. Natuurlijk wil ik Tjaard, mijn kamergenoot bedanken. Je bent hét klankbord voor allerlei lab-issues, vooral PC-problemen. Bedankt voor je steun in slechte en goede tijden. Anita, bedankt dat je mijn paranimf wilt zijn. Op één of andere manier komen we elkaar steeds weer tegen zowel werk gerelateerd als privé.

Ten slotte wil ik mijn familie en vrienden bedanken. Bedankt lieve dames van het aquarobic clubje van de woensdagavond. Zullen we nog even met de slurven meppen? Ook wil ik de vrienden van de Lotustours bedanken voor hun belangstelling.

Pap en Mam bedankt voor de goede zorgen en steun door je jaren heen. Helaas mag Mam het niet meer meemaken maar ze was altijd mijn toeverlaat. Jakob, bedankt dat je mijn paranimf bent. Mijn kinderen Lisa en Bo Wan bedank ik, gewoon omdat jullie er zijn. Nu word ik eindelijk een soort dokter ‘in de kristallen’. Blijven jullie nog even in “Hotel papa- mama”? Cees, mijn rots in de branding, mijn laatste woorden zijn voor jou. Bedankt voor de steun en de relativerende woorden die mij soms weer met beide benen op de grond hebben gezet.

September 2014

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