Insights Into the Catalytic Mechanisms of the Protein Enzymes, PFKFB3

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2012 Insights into the catalytic mechanisms of the protein enzymes, PFKFB3 and VldE, using x-ray crystallography Michael Christopher Cavalier Louisiana State University and Agricultural and Mechanical College

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INSIGHTS INTO THE CATALYTIC MECHANISMS OF THE PROTEIN ENZYMES, PFKFB3 AND VLDE, USING X-RAY CRYSTALLOGRAPHY

A Dissertation

Submitted to the Graduate Faculty of the Louisiana State University and Agricultural and Mechanical College in partial fulfillment of the requirements for the degree of Doctor of Philosophy

The Department of Biological Sciences

by Michael Christopher Cavalier B.S., Louisiana State University, 2006 December 2012

ACKNOWLEDGMENTS

I would like to begin by thanking my major professor, Dr. Yong-Hwan Lee, for taking me into his laboratory and providing me with the opportunity to learn the skill of protein crystallography. I thank Dr. Marcia Newcomer, Dr. Grover Waldrop, and Dr. Bing-Hao Luo for joining my graduate committee and for all the advice they have offered. I thank Dr. Henry

Bellamy and Dr. David Neau for contributing their expertise in synchrotron data collection. I thank Dr. Erin Hawkins for her guidance during my teaching duties and for her continuing encouragement. I thank my fellow lab members; Blake Crochet, Dr. Jeong-Do Kim, Dr. Young-

Sun Yim, Dr. Song-Gun Kim, and Minsuh Seo for the valuable discussions and for the contributions they have made to my education.

I also offer a special thanks to my parents, Bonnie and Nickie, and to my brother,

Matthew, for their unrelenting support during this long trying process. I also thank my fiancée,

Heather, whose patient love enabled me to complete this work. This would not be possible without them.

TABLE OF CONTENTS

ACKNOWLEDGMENTS ...... ii

LIST OF TABLES ...... vi

LIST OF FIGURES ...... vii

ABSTRACT ...... ix

CHAPTER 1: REVIEW OF LITERATURE, THE BIFUNCTIONAL ENZYME ...... 1 Carbohydrate Metabolism ...... 1 History of Metabolism ...... 1 Glycolysis ...... 1 Gluconeogenesis ...... 4 The Discovery of Fructose-2,6-Bisphosphate and the Bifunctional Enzyme ...... 5 Two Functional Folds ...... 8 Opposing Reactions ...... 8 Histidine Phosphatases...... 8 P-loop Kinases ...... 11 The Tissue Specific Isoforms of the Bifunctional Enzyme ...... 14 Conserved Structure ...... 14 Liver ...... 18 Heart ...... 19 Testis ...... 19 Inducible ...... 20 PFKFB3 and the Warburg Effect ...... 20 The Glycolytic Phenotype of Cancer ...... 20 PFKFB3 and Cancer ...... 21 The Hypoxia-Inducible Transcription Factor HIF-1...... 22 PFKFB3 as a Therapeutic Target ...... 23

CHAPTER 2: MOLECULAR BASIS OF THE FRUCTOSE-2,6- BISPHOSPHATASE REACTION OF PFKFB3: TRANSITION STATE AND THE C-TERMINAL FUNCTION ...... 25 Introduction ...... 25 Results ...... 29 Overall Structure ...... 29 PFKFB3-P•F-6-P Structure ...... 29 PFKFB3•AlF4 Structure ...... 30 PFKFB3•PPi Structure ...... 33 Comparison of Bisphosphatase Active Sites ...... 33 Degradation of Fructose-2,6-Bisphosphate ...... 36 Discussion ...... 40 Materials and Methods ...... 42 Preparation and Crystallization of PFKFB3 ...... 42

iii

Data Collection and Processing ...... 43 Structure Determination and Refinement ...... 43

CHAPTER 3: REVIEW OF LITERATURE, VLDE ...... 46 Diversification of Natural Products ...... 46 Natural Products Offer Unique Scaffolding ...... 46 Diversification of the Erythromycin Scaffold by Genetic Manipulation...... 47 Diversification of the Avermectin Scaffold by Alternative Precursors ...... 50 Diversification of the Antitumor Indolocarbazoles by Recombinant Enzymes ....51 The Aminocyclitol class of Natural Products ...... 52 The Variety of Aminocyclitols ...... 52 The Aminoglycosides ...... 53 The Five-Membered Ring Aminocyclitols ...... 56 The C7N Aminocyclitols ...... 58 Validamycin A ...... 61 Early Uses of Validamycin A ...... 61 Discovery of the Validamycin A Gene Cluster ...... 62 Biosynthetic Pathway of Validamycin A ...... 62 Glycosyltransferases ...... 66 A Large and Diverse Class of Enzymes ...... 66 GT-A and GT-B Topologies ...... 67 Inverting Glycosyltransferases ...... 69 Retaining Glycosyltransferases ...... 72 Oxocarbenium Ion-Like Transition State ...... 74

CHAPTER 4: MECHANISTIC INSIGHTS INTO VALIDOXYLAMINE A 7'-PHOSPHATE SYNTHESIS BY VLDE USING THE STRUCTURE OF THE ENTIRE PRODUCT COMPLEX ...... 75 Introduction ...... 75 Materials and Methods ...... 80 Purification and Crystallization ...... 80 Data Collection and Processing ...... 84 Structure Determination and Refinement ...... 85 Results ...... 89 Overall Structure ...... 89 The Nucleotide Binding Site ...... 91 The Cyclitol Binding Sites ...... 93 Observed Changes in Conformations ...... 97 Discussion ...... 101 Mechanistic Considerations ...... 101 Significance...... 102

CHAPTER 5: CONCLUSIONS ...... 105

REFERENCES ...... 107

APPENDIXES ...... 131 A. A Crystallographic Study ...... 131 B. Copyright Permissions ...... 132

VITA ...... 138

LIST OF TABLES

Table 2.1 Notable interactions between the bisphosphatase active site ligands and the protein ...... 31

Table 2.2 Statistics of reflection data and structure refinements (PFKFB3) ...... 44

Table 4.1 Statistics of reflection data and structure refinements (VldE) ...... 86

Table 4.2 Notable interactions between VldE and ligands ...... 94

Table 4.3 DALI server results ...... 103

LIST OF FIGURES

Figure 1.1 Glycolysis and Gluconeogenesis ...... 3

Figure 1.2 Fructose-2,6-Bisphosphate...... 7

Figure 1.3 The catalytic mechanism of histidine phosphatases ...... 10

Figure 1.4 P-loop NTPases ...... 12

Figure 1.5 The crystal structures of the bifunctional enzyme ...... 15

Figure 1.6 Covalent modification of the bifunctional enzyme ...... 16

Figure 1.7 The kinase reaction ...... 17

Figure 2.1 Overall folding of PFKFB3 and sites of ligand binding ...... 26

Figure 2.2 Sequence alignment of PFKFB ...... 27

Figure 2.3 The minimal enzymatic reaction ...... 30

Figure 2.4 Aluminum tetrafluoride within the active site ...... 32

Figure 2.5 Pyrophosphate within the active site...... 34

Figure 2.6 A hypothetical representation of the Michaelis complex (E•F-2,6-P2) ...... 37

Figure 2.7 The complete mechanism of phospho-histidine formation/degradation ...... 40

Figure 3.1 Differentiated Scaffolds ...... 48

Figure 3.2 The aminoglycosides ...... 54

Figure 3.3 The aminocyclopentitols ...... 57

Figure 3.4 The C7N aminocyclitols ...... 59

Figure 3.5 Early validamycin biosynthesis ...... 63

Figure 3.6 Late validamycin biosynthesis ...... 65

Figure 3.7 Glycosyltransferase topologies ...... 68

Figure 3.8 Catalytic mechanisms of glycosyltransferases...... 70

vii

Figure 4.1 The VldE and OtsA reactions ...... 76

Figure 4.2 An alignment of VldE and OtsA ...... 78

Figure 4.3 The overall fold of VldE ...... 80

Figure 4.4 The catalytic site ...... 81

Figure 4.5 Ligand interactions...... 92

Figure 4.6 Trehalose within the catalytic site...... 97

Figure 4.7 Conformational changes ...... 98

Figure 4.8 The SNi mechanism of VldE ...... 103

viii

ABSTRACT

This work describes the crystallographic studies of two enzymes and provides mechanistic insights into their respective catalytic processes. The first study investigates the molecular basis of the fructose-2,6-bisphosphatase reaction of the inducible isoform of the bifunctional enzyme, 6-phosphofructo-2-kinase/fructose-2,6-bisphosphatase (PFKFB3). The bifunctional enzyme is solely responsible for the cellular concentration of a regulator of glucose metabolism, fructose-2,6-bisphosphate. PFKB3 was investigated using the crystal structures of the enzyme in a phospho-enzyme intermediate state (PFKFB3-P•F-6-P), in a transition state- analogous complex (PFKFB3•AlF4), and in a complex with pyrophosphate (PFKFB3•PPi). With these structures, the structures of the Michaelis complex and the transition state were extrapolated. Additionally the C-terminal domain (residues 440-446) was rearranged in

PFKFB3•PPi, implying that this domain plays a critical role in binding of substrate to, and release of product from, the bisphosphatase catalytic pocket. These findings provide a new insight into the understanding of the phosphoryl transfer reaction.

The second study investigates the molecular basis of the reaction catalyzed by the pseudo-glycosyltransferase, VldE. VldE catalyzes non-glycosidic C-N coupling between an unsaturated cyclitol and a saturated aminocyclitol with the conservation of the anomeric configuration to form validoxylamine A 7´-phosphate, the biosynthetic precursor of the antibiotic validamycin A. To study the molecular basis of its mechanism, the three-dimensional structures of VldE from Streptomyces hygroscopicus subsp. limoneus was determined in apo form, in complex with GDP, in complex with GDP and validoxylamine A 7´-phosphate, and in complex with GDP and trehalose. The structures of VldE with the catalytic site in both an “open” and

“closed” conformation are also described. With these structures, the preferred binding of the

ix guanine moiety by VldE, rather than the uracil moiety as seen in OtsA, could be explained. The elucidation of the VldE structure in complex with the entirety of its products provides insight into the internal return mechanism by which catalysis occurs with a net retention of the anomeric configuration of the donated cyclitol.

CHAPTER 1: REVIEW OF LITERATURE, THE BIFUNCTIONAL ENZYME

Carbohydrate Metabolism

History of Metabolism

The term “metabolism”, which is derived from the Greek word “metabolē” for “change” is defined as “the chemical changes in living cells by which energy is provided for vital processes and activities and new material is assimilated” (Merriam-Webster Inc. 2005). In the early 20th century, the term “metabolism” was synonymous with the term “nutrition” as both are centered around the intake of nutrients as a sources of usable energy and precursors of biological compounds (Bing 1971).

The scientific study of human metabolism is generally credited to originate with the

Italian physician Santorio Santorio. As recounted in his 1614 book Ars de Statica Medicina,

Santorio performed self-experimental studies on what he termed “insensible perspiration”

(Eknoyan 1999). Not until the 19th century works of Louis Pasteur, who demonstrated that the fermentation of sugar by yeast was a biological process within its life cycle, and Friedrich

Wöhler, whose chemical synthesis of urea demonstrated that biological compounds could be produced from non-organic chemicals, was it understood that the transformation of nutrients into energy and biological compounds was not different from the general processes of chemistry

(Dubos 1951; Kinne-Saffran and Kinne 1999). With the discovery of enzymes by Eduard

Buchner at the beginning of the 20th century(Jaenicke 2007), biochemistry was born and metabolism could now be thought of as a series of chemi-enzymatic reactions.

Glycolysis

Metabolism can be divided into two sets of processes, the catabolic processes in which organic compounds are broken-down and the anabolic processes in which cellular components

1 are synthesized. Glycolysis, also known as the Embden-Meyerhoff-Parnas pathway, is a catabolic pathway through which there is an enzymatic degradation of carbohydrates into pyruvate while energy is harvested in the forms of adenosine triphosphate (ATP) and reduced nicotinamide adenine dinucleotide (NADH). In an aerobic environment, pyruvate is further degraded into carbon dioxide and water by the citric acid cycle in order to send reducing equivalents into oxidative phosphorylation. In human cells, the presence of an anaerobic environment will cause the fermentation of pyruvate into lactic acid in order to regenerate oxidized nicotinamide adenine dinucleotide (NAD+), which would allow the continuation of glycolysis.

Although the glycolytic degradation of a single molecule of the carbohydrate glucose into two pyruvates is mediated by a pathway of ten enzymes, three of these enzymatic reactions are under a particularly stringent regulation (Figure 1.1). Because of the investment of a high energy storage compound to the reaction in the form of ATP, the conversion of glucose to glucose-6-phosphate by hexokinase is tightly regulated. Likewise, the investment of an ATP in the conversion of fructose-6-phosphate (F6P) to fructose-1,6-bisphosphate (F-1,6-P2) by 6- phosphofructose-1-kinase (PFK-1) gives need for tight regulation. The investment of second

ATP is a committal step and following this further investment glycolysis must proceed to completion. The final step of glycolysis, the generation of pyruvate from phosphoenolpyruvate

(PEP) by pyruvate kinase, is also strictly controlled. Although the generation of glycolytic products and product intermediates allow for cells to meet their energy demands while also producing biosynthetic precursors for a series of biological compounds, glycolysis must be regulated in order to balance the various metabolic processes within the cell. (McKee and

McKee 2009)

Figure 1.1. Glycolysis and Gluconeogenesis. Shown are the key points of regulation of glycolysis and gluconeogenesis within glucose metabolism.

Gluconeogenesis

The liver is responsible for the maintenance of blood glucose levels so that other tissues such as muscle or brain may meet their individual energy demands. To fulfill this responsibility, liver cells must be capable of regenerating glucose so that it can be introduced back into the blood stream and transported to the various energy requiring tissues. One such method would be the reformation of glucose by the enzymatic pathway known as gluconeogenesis.

Gluconeogenesis can generate glucose from metabolic precursors within the liver cells such as pyruvate, or from lactate taken in from the blood stream via the Cori Cycle. Lactate can be easily converted back to pyruvate by the enzyme lactate dehydrogenase.

Although similar to a reversal of the glycolytic pathway, gluconeogenesis must bypass the three tightly regulated and irreversible enzymatic reactions of glycolysis through an alternative set of enzymes (Figure 1.1). Pyruvate must be carboxylated by pyruvate carboxylase before being decarboxlyated and phosphorylated by PEP carboxykinase during the regeneration of phosphoenolpyruvate. Phosphoenolpyruvate will then be passed through a series of reversed glycolytic reactions until fructose-1,6-bisphosphate is regenerated. The phosphofructose-1- kinase reaction is again irreversible, and the dephosphorylated product, fructose-6-phosphate, must be generated by fructose-1,6-bisphosphatase (F-1,6-P2ase). Gluconeogenesis would again proceed through a reverse of glycolysis until glucose is regenerated by the dephosphorylation of glucose-6-phosphate by glucose-6-phosphatase. Upon the completion of gluconeogenesis, glucose is now available for release into the blood stream by the liver cells. Like the corresponding reactions within glycolysis, these gluconeogenic reactions are under stringent regulation, typically in a manner that is reciprocal to the regulation of glycolysis (McKee and

McKee 2009).

The Discovery of Fructose-2,6-Bisphosphate and the Bifunctional Enzyme

As previously mentioned, phosphofructose-1-kinase (PFK-1) is an extremely important regulatory enzyme within glycolysis. PFK-1 catalyzes the production of fructose-1,6- bisphosphate from ATP and fructose-6-phosphate. The reaction, which is made important by the investment of the second ATP to glycolysis, is considered the first committedstep to glycolysis.

Curiously, it was shown that at nearly physiological conditions including physiological concentrations of its substrates and of its known effectors (F-1,6-P2, adenosine monophosphate

(AMP), and orthophosphate (Pi)), purified PFK-1 is nearly inactive in vitro (Reinhart and Lardy

1980). Because purified PFK-1 showed ample activity and because the activity of PFK-1 in physiological conditions could be restored by reintroducing the contents of the low molecular weight fractions obtained during the purification process, researchers realized that there must be an additional activating factor lost during purification (Furuya and Uyeda 1980; Van

Schaftingen, Hue et al. 1980). Further investigation determined that this new low molecular weight effector, whose cellular concentration is reduced by the introduction of glucagon or epinephrine, works by relieving the inhibition of PFK-1 caused by the binding of a second ATP molecule to an allosteric site on PFK-1 at cellular concentrations of ATP (Furuya and Uyeda

1980; Richards and Uyeda 1980).

Many preliminary observations regarding this stimulator were made. The cellular concentration of the stimulator was increased by the introduction of glucose, and had a molecular weight similar to F-1,6-P2. Because the stimulator was acid-labile and was capable of being destroyed by alkaline phosphatase, it was assumed that the stimulator must be a phosphoric ester

(Van Schaftingen, Hue et al. 1980). Further analysis of the products generated by acid driven hydrolysis showed that the stimulator consisted solely of F-6-P and Pi. Researchers suggested

5 that the stimulator was most probably fructose-2,6-bisphosphate (F-2,6-P2) (Van Schaftingen,

Hue et al. 1980). This was confirmed using synthetic F-2,6-P2 to reproduce stimulation, and the anomeric form of the stimulator was determined to be -D-fructose-2,6-bisphosphate (Figure

1.2A) (Van Schaftingen and Hers 1980; Furuya, Uyeda et al. 1981; Uyeda, Furuya et al. 1981).

With the discovery of a small molecular weight stimulator of a rate-limiting enzyme within glycolysis, it was only natural to investigate what effect, if any, the compound had on gluconeogenesis. Because PFK-1 and fructose-1,6-bisphosphatatase (F-1,6-P2ase) were already known to share many effectors in a reciprocal manner, it was thought that F-1,6-P2ase was a likely target of F-2,6-P2. Indeed, F-2,6-P2 was shown to be capable of inhibiting the enzymatic activity of this key regulatory enzyme of gluconeogenesis (Pilkis, El-Maghrabi et al. 1981). As both PFK-1 and F-1,6-P2ase would otherwise be involved in a futile cycle between F-1,6-P2 synthesis and degradation, discovery of F-2,6-P2 expanded the understanding of the cellular coordination of glycolysis and gluconeogenesis (Uyeda, Furuya et al. 1982).

In 1981, Furuya and Uyeda released a publication in which they detailed the discovery of a liver enzyme capable of synthesizing the PFK-1 stimulator, F-2,6-P2. This enzyme, 6- phosphofructo-2-kinase (6-PF-2-Kase), catalyzes the reaction between D-fructose-6-phosphate and ATP in the presence of magnesium in order to form D-fructose-2,6-bisphosphate and adenosine diphosphate (ADP) (Figure 1.2B) (Furuya and Uyeda 1981). It was determined that the activity of 6-PF-2-Kase was sensitive to covalent modification by phosphorylation.

Specifically, the enzyme was inactivated by phosphorylation by cyclic-AMP dependent kinase and activated by dephosphorylation by phosphatase (Van Schaftingen, Davies et al. 1981;

Furuya, Yokoyama et al. 1982).

Figure 1.2. Fructose-2,6-Bisphosphate. Shown is (A) the chemical structure of fructose-2,6- bisphosphate, an effector of glycolytic flux, and (B) the enzymatic reactions responsible for the synthesis and degradation of this effector.

Within a year of the discovery of 6-PF-2-Kase, an enzyme responsible for the degradation of F-2,6-P2 into F-6-P and Pi was also found within liver extracts (Figure 1.2B) (El- maghrabi, Claus et al. 1982; Furuya, Yokoyama et al. 1982). Inversely to 6-PF-2-Kase, the hydrolytic activity of this new enzyme, fructose-2,6-bisphosphatase (F-2,6-P2ase), was activated by cyclic-AMP dependent kinase and inactivated by glucagon and epinephrine (Furuya,

Yokoyama et al. 1982; Richards, Yokoyama et al. 1982). Strangely, 6-PF-2-Kase and F-2,6-

P2ase coeluted from phosphocellulose columns, were of similar molecular weights, and consisted of similarly sized catalytic units. It did not take long for researchers to realize that these two

7 enzymes were in fact a single bifunctional enzyme with opposing catalytic functions and reciprocal methods of regulation (El-maghrabi, Claus et al. 1982; Pilkis, Chrisman et al. 1983).

Two Functional Folds

Opposing Reactions

The bifunctional enzyme, 6-phosphofructo-2-kinase/fructose-2,6-bisphosphatase, is solely responsible for the cellular concentration of the glycolytic regulator, fructose-2,6- bisphosphate. It fulfills its role by concertedly synthesizing and degrading this effector.

Although this single polypeptide catalyzes both opposing reactions, the reactions do not necessarily proceed through analogous mechanisms (Kim, Cavalier et al. 2007). The phosphotransferase activity of the N-terminal domain, which uses ATP as a source for the high- energy phosphoester bond, is mediated by a fold belonging to the P-loop NTPase superfamily

(Hasemann, Istvan et al. 1996; Leipe, Koonin et al. 2003). Conversely, the hydrolytic activity of the C-terminal domain is mediated by a fold belonging to the histidine phosphatase superfamily which does not regenerate ATP but instead yields orthophosphate (Hasemann, Istvan et al. 1996;

Rigden 2008).

Histidine Phosphatases

The histidine phosphatase superfamily can be divided into two evolutionarily related branches, the phosphoglycerate mutase family which includes fructose-2,6-bisphosphatase and the acid phosphatase family which also includes the phytases (Rigden 2008). Although the highest observed percentage identity between the two branches of enzymes is only 18%, there is an unambiguous evolutionary relationship between the sequences of both branch members as was further illustrated upon the solving of the first crystal structure of the first acid phosphatase family enzyme in 1993 (Schneider, Lindqvist et al. 1993; Rigden 2008). Members of both

8 branches share a RHG motif near the beginning of the fold, whose histidine is known to be phosphorylated during catalysis (Bazan, Fletterick et al. 1989). It should be noted that there is a clear distinction between the subcellular localization of the enzyme members of the two branches. While the phosphoglycerate mutase family enzymes are generally found in the cytoplasm or the nucleus, acid phosphatase enzymes enter into the secretory pathway and are found in the endoplasmic reticulum, the cell surface, periplasm, cell wall, or secretions (Rigden

2008). The Pfam database (http://pfam.sanger.ac.uk/) classifies over 15,000 sequences within the phosphoglycerate mutase family and over 2,600 sequences within the acid phosphatase family with both families having extensive prokaryotic and eukaryotic representatives.

However, there are no acid phosphatase family members and only 86 phosphoglycerate mutase family members found within archaea (as of June 16, 2012) (Punta, Coggill et al. 2012).

The histidine phosphatases are diverse in function and are known to play roles in many metabolic pathways. Examples would include the cofactor-dependent phosphoglycerate mutase

(dPGM) within glycolysis/gluconeogenesis, adenosylcobalamin-5´-P phosphatase (CobC) in vitamin B12 biosynthesis (Zayas and Escalante-Semerena 2007), 5´- phosphoribostamycinphosphatase (NeoP) within neomycin biosynthesis (Huang, Haydock et al.

2005; Kudo and Eguchi 2009), and the fungal phosphate scavenging enzymes such as nucleotide phosphatase (Pho5p) and phytase (Irving and Cosgrove 1972; Kennedy, Pillus et al. 2005).

Histidine phosphatases are also known to play roles in signaling/regulation. For example, fructose-2,6-bisphosphatase degrades a small molecule responsible for the regulation of glycolytic flux while the phosphomutase (PMU1) regulates the concentration of a small molecule influencing transcription of adenine synthesis genes in yeast, and the eukaryotic multiple inositol

9 polyphosphate phosphatase 1(Minpp1) regulates the concentrations of other signaling molecules

(Chi, Yang et al. 2000; Rebora, Laloo et al. 2005; Michels and Rigden 2006).

The catalytic mechanism of the histidine phosphatases works through the phosphorylation of a histidine residue to form a phospho-enzyme intermediate state followed by its dephosphorylation (Figure 1.3). Neighboring residues, in particular a second histidine and two arginines form a phosphate pocket and stay in contact with the phosphate oxygens before during and after the linear transfer of phosphate. The conserved RHG motif of histidine phosphatases is completed by a glycine residue (sometimes alanine) which packs closely to the protein core and whose carbonyl makes an important hydrogen bond with the phosphorylated histidine side-chain placing the group in an appropriate position for catalysis. There is also a conserved L[S/T]XXG motif in the second layer behind the catalytic site which adds to the maintenance of the histidine conformation which is extremely important for catalysis (Rigden

2008).

Figure 1.3. The catalytic mechanism of histidine phosphatases. Shown is the prototypical histidine phosphatase catalyzed reaction with a phospho-histidine intermediate step.

P-loop Kinases

Found within prokaryotes, eukaryotes and archaea, the P-loop NTPase superfamily is the most abundant and diverse of several distinct nucleotide triphosphate (NTP)-binding folds and was estimated to comprise 10-18% of the predicted gene products of the sequenced prokaryotic and eukaryotic genomes (Koonin, Wolf et al. 2000; Leipe, Koonin et al. 2003). The P-loop

NTPases are characterized by Walker A (P-loop) and B motifs responsible for the binding of the

and  phosphates of the NTP as well as a Mg2+ cation (Figure 1.4). The Walker A motif adopts a GXXXXGK[ST] sequence pattern while the Walker B motif (hhhhD, where h is a hydrophobic residue) contains a conserved aspartate (occasionally glutamate) which coordinates the Mg2+ with the  and  phosphates. This conserved aspartate also interacts with the conserved serine/threonine of the Walker A motif which properly positions the two phosphate-binding domains (Walker, Saraste et al. 1982).

Amongst the P-loop NTPases are the P-loop kinases which transfer the -phosphate of the

NTP to a wide range of substrates. Outside of P-loop folds, kinases have evolved independently within a number of folds such as Rossman, RRM-like, ribonuclease H, and TIM / barrel folds

(Cheek, Zhang et al. 2002). Amongst the best characterized P-loop kinases are the nucleotide kinases such as adenylate kinase (AK) and the small molecule kinases such as 6-phosphofructo-

2-kinase and shikimate kinase. Shikimate kinase transfers the terminal phosphate of ATP to the

3-hydroxyl of shikimic acid and is part of the biosynthesis of chorismic acid, a precursor for aromatic compounds essential to plants and micro-organisms (Krell, Coggins et al. 1998). Of course, 6-phosphofructo-2-kinase is responsible for the synthesis of a regulator of glucose metabolism, fructose-2,6-bisphosphate (Kim, Cavalier et al. 2007). Adenylate kinase catalyzes

Figure 1.4. P-loop NTPases. Shown is the typical binding of a nucleotide triphosphate by the Walker A and B motifs of P-loop NTPases.

12 the transfer of a phosphate from ATP to AMP which links AMP to the ADP/ATP equilibrium of the cell (Muller and Schulz 1992).

Amongst the P-loop NTPases, the P-loop kinases are most similar to the GTPases which include the enzymes Ras and the elongation factor Tu (EF-Tu) (Leipe, Wolf et al. 2002; Leipe,

Koonin et al. 2003). However, P-loop kinases are clearly discernible by the presence of a mostly helical LID module. The LID module contributes an arginine residue (asparagine in the case of

6-PF-2-Kase), which interacts with the nucleotide base, as well as a second conserved basic residue (arginine or lysine), which is thought to stabilize the transition state by neutralizing the developing negative charge on the - oxygen atom (Leipe, Koonin et al. 2003).

Upon the elucidation of the crystallographic structure of 6-PF-2-Kase in 1996, a structural relationship between 6-PF-2-Kase, the nucleotide kinases, and the GTPases was immediately apparent (Hasemann, Istvan et al. 1996). The structural similarity between the catalytic sites of 6-PF-2-Kase and adenylate kinase supported the recent rejection of a proposed

6-PF-2-Kase mechanism of catalysis operating through a general base which would activate the sugar hydroxyl group to potentiate its nucleophilic attack of the -phosphate of ATP. This rejected mechanism was based on the assumption that the phosphorylation of the fructose-6- phosphate at the 2-hydroxyl group by 6-PF-2-Kase would proceed through a mechanism similar to the phosphorylation of the 1-hydroxyl group by 6-phosphofructose-1-kinase. 6-PF-2-Kase, like adenylate kinase, lacks such a nucleophile. Instead, 6-PF-2-Kase is thought to only mediate a transition state stabilization (Hasemann, Istvan et al. 1996). The P-loop kinases do not contain a traceable ancestral nucleophile, although individual P-loop kinases have independently acquired one on multiple occasions (Leipe, Koonin et al. 2003).

The Tissue Specific Isoforms of the Bifunctional Enzyme

Conserved Structure

In 1996, the first structural characterization of the bifunctional enzyme, in its entirety, was performed using x-ray crystallographic techniques on the testis isoenzyme (Hasemann,

Istvan et al. 1996). Since, the crystallographic structures of the liver and inducible isoforms have also been elucidated (Lee, Li et al. 2003; Kim, Manes et al. 2006). The biologically active unit of the bifunctional enzyme is a homodimer (Figure 1.5). Each monomer consists of two clearly divided catalytic domains. The N-terminal half of the polypeptide forms the kinase functional domain, while the C-terminal half forms the bisphosphatase functional domain (Hasemann,

Istvan et al. 1996). All tissue-specific isoforms of the bifunctional enzyme, whose structures have been elucidated, tightly conserve the fold of the two catalytic domains. The main points of divergence are the C- and N- terminal segments (Hasemann, Istvan et al. 1996; Lee, Li et al.

2003; Kim, Manes et al. 2006). As described elsewhere within this document, these terminal segments encode phosphorylation sites which allow for regulation of the cellular concentrations of F-2,6-P2 by fluctuating the balance of kinase and bisphosphatase activities (Figure 1.6).

The N-terminal catalytic domain (6-PF-2-K) consists of a six stranded -sheet surrounded by seven-helices. Within this fold, the binding of ATP is facilitated by a Walker A motif (Gly-X-X-Gly-X-Gly-Lys-Thr). The adenosine moiety is stacked within non-polar residues. A Walker B motif (Z-Z-Z-Z-Asp, where Z is any hydrophobic residue) orients the octahedral Mg2+ ion coordination between the protein and the - and -phosphates of ATP

(Hasemann, Istvan et al. 1996). These motifs are commonly associated with the adenylate kinase

(AK), Ras, and EF-Tu families of nucleotide binding proteins (Walker, Saraste et al. 1982; Traut

1994). F-6-P and ATP binding loops conform around the substrates to exclude water from the

Figure 1.5. The crystal structures of the bifunctional enzyme. Shown are ribbon diagrams representing the crystal structures of the tissue-specific isoforms of 6-phosphofructo-2- kinase/fructose-2,6-bisphosphatase.

Figure 1.6. Covalent modification of the bifunctional enzyme. Shown are the phosphorylation sites unique to each tissue-specific isoform of 6-phosphofructo-2- kinase/fructose-2,6-bisphosphatase. active site (Hasemann, Istvan et al. 1996). The substrates ATP and F-6-P are oriented so that there is a strong direct substrate-substrate interaction between the -phosphate of ATP and the

O2 oxygen of F-6-P. Upon the substrate-assisted deprotonation of the O2 oxygen, F-6-P can perform a nucleophilic attack on the -phosphate of ATP (Kim, Cavalier et al. 2007). This is followed by an “in-line” transfer of phosphate accompanied with the inversion of the phosphate stereochemistry (Kitajima, Sakakibara et al. 1984; Lahiri, Zhang et al. 2003; Klahn, Rosta et al.

2006; Wittinghofer 2006). The Mg2+ ion bound by the Walker B motif holds the-phosphate of

ATP in the proper position for catalysis (Figure 1.7) (Vertommen, Bertrand et al. 1996).

The C-terminal catalytic domain (F-2,6-P2ase) also consists of a central -sheet with surrounding -helices. An additional subdomain of two helices contributes residues to substrate binding (Hasemann, Istvan et al. 1996). F-2,6-P2ase belongs to the acid phosphatases and

Figure 1. 7. The kinase reaction. Shown is reaction catalyzing the synthesis of fructose-2,6- bisphosphate by the kinase domain of phosphofructo-2-kinase/fructose-2,6-bisphosphatase. phosphoglycerate mutase family of enzymes (Bodansky 1972; Fothergillgilmore and Watson

1989). These enzymes worked through a shared mechanism involving a covalent phosphor- histidine intermediate within a conserved sequence motif (Z-X-Arg-His-Gly- Glu/Gln-X-X-X-

Asn, where Z is a hydrophobic residue) (Rose 1970; Vanetten and Hickey 1977; Pilkis,

Walderhaug et al. 1983).

The active homodimer assembles in a head-to-head manner. The N-terminal kinase domains interact in a manner by which the six stranded -sheet of each is oriented to form a single twelve stranded -sheet. The kinase domains scarcely interact at all (Hasemann, Istvan et al. 1996).

Liver

To account for the metabolic demands of the individual tissues, there are four tissue- specific isoforms of the bifunctional enzyme (also known as phosphofructo-2-kinase/fructose-

2,6-bisphosphatase or PFKFB). The first of which is the liver isoform (PFKFB1), which is encoded by the gene, pfkfb1 (Algaier and Uyeda 1988). A splice variant of the pfkfb1 gene is also responsible for the encoding of the skeletal form of the bifunctional enzyme, which has an alternative N-terminus (Darville, Crepin et al. 1989).

PFKFB1 contains one phosphorylation site near the N-terminus (Figure 1.7). In response to glucagon, cAMP-dependent protein kinase (PKA) is stimulated, and PFKFB1is phosphorylated at Ser32. (Murray, Elmaghrabi et al. 1984). This results in an inactivation of the

6-PF2-K activity in concert with an activation of the F-2,6-P2ase activity (Van Schaftingen,

Davies et al. 1981; El-maghrabi, Claus et al. 1982; Van Schaftingen, Davies et al. 1982;

Sakakibara, Kitajima et al. 1984). The kinase:bisphosphatase ratio of catalysis shifts from 1.4 to

0.2, a shift from a predominant F-2,6-P2 synthesis to a predominant F-2,6-P2 degradation (El-

Maghrabi, Noto et al. 2001). The resulting decrease in F-2,6-P2 concentration will slow glycolysis while stimulating gluconeogenesis within the liver. Because of the alternative splicing, the skeletal isoform lacks this phosphorylation site, and its activity is not subject to phosphorylation by PKA (Crepin, Darville et al. 1989). By comparing the skeletal enzyme to the liver form, the role of the N-terminus is regulation becomes even more apparent. The

18 skeletal enzyme has a kinase:bisphosphatase catalytic ratio of only 0.04, which is analogous to the net direction of F-2-6-P2 degradation seen in the phosphorylated liver isoform. As skeletal tissue is highly glycolytic, the isoform is well suited to the metabolism of the tissue. The lack of a regulatory site leaves the net direction of catalysis by the skeletal form solely dependent of the concentration of F-6-P (El-Maghrabi, Noto et al. 2001).

Heart

In 1985, the gene product of pfkfb2 was discovered (Rider, Foret et al. 1985). The heart isoform (PFKFB2) shares highly conserved catalytic domains with the liver isoform. The primary differences between the isoforms are at the C- and N-termini (Figure 1.7) (Darville,

Chikri et al. 1991). PFKFB2 does not contain a single phosphorylation site on the N-terminus but instead two sites on the C-terminus. In opposition to PFKFB1, phosphorylation of PFKFB2 at these sites by PKA and insulin-stimulated protein kinase leads to an activation of kinase activity and an inactivation of the bisphosphatase activity (Deprez, Vertommen et al. 1997).

Testes

In 1991, the gene product of pfkfb4 was cloned and characterized (Sakata, Abe et al.

1991). The testes isoform (PFKFB4) also shares highly conserved catalytic domains with the liver and isoform. Again, it is the C- and N-termini which diverge from the other isoforms. Like the skeletal isoenzyme, PFKFB4 does not contain any sites of phosphorylation and is not subject to regulation by PKA (Figure 1.7). Testes tissues are also highly glycolytic and the kinetics of the testes isoform of the bifunctional enzyme suggest that is an intermediate between the muscle and liver forms which has a higher kinase:bisphosphatase ratio and is active at lower substrate concentrations (El-Maghrabi, Noto et al. 2001).

Inducible

The gene product of pfkfb3 is a PFKFB isoenzyme (PFKFB3) that is ubiquitously expressed (Manzano, Rosa et al. 1998). PFKFB3 is the same isoenzyme that been previously reported to be contained within the tissues of the brain, placenta, and pancreatic -islets.

(Ventura, Rosa et al. 1992; Sakakibara, Okudaira et al. 1999; El-Maghrabi, Noto et al. 2001). As

PFKFB3 expression is stimulated by hypoxia, it is often referred to as the inducible isoform

(Minchenko, Leshchinsky et al. 2002). PFKFB3, which has the highest kinase:bisphosphatase activity ratio, allows for elevated F-2,6-P2 levels, and thus an increase in glycolytic flux. In fact, the bisphosphatase domain of the inducible isoform is nearly inactive (Sakakibara, Okudaira et al. 1999). PFKFB3 has been linked to the elevated glycolytic activity within cancer cells

(Chesney, Mitchell et al. 1999). The inducible isoform does contain a single site for phosphorylation by PKA (Figure 1.7). Phosphorylation further elevates the kinase:bisphosphatase activity ratio of PFKFB3 (Sakakibara, Kato et al. 1997). The 2006 structure of PFKFB3 described the N-terminus exists as a -hairpin motif which makes contact with the bisphosphatase domain (Figure 1.5B) (Kim, Manes et al. 2006). This can help to explain the regulatory effect of the N-terminus of the C-terminal bisphosphatase domain and vice versa.

PFKFB3 and the Warburg Effect

The Glycolytic Phenotype of Cancer

In order to proceed through the processes of uncontrolled cell division and proliferation, cancer cells uptake glucose at levels that are in far excess of what is needed to meet energetic demands (Bauer, Harris et al. 2004; DeBerardinis, Lum et al. 2008). Curiously, cancer cells continue with a high level of glucose uptake and glycolysis despite the availability of O2 (aerobic

20 glycolysis) (Gatenby and Gillies 2004; Moreno-Sanchez, Rodriguez-Enriquez et al. 2007). This phenomena was first described by Dr. Otto Warburg in 1956, and thus it is often recognized as the “Warburg Effect” (Warburg 1956). The rapid uptake of glucose is the basis by which the glucose analog, 18F-fluorodeoxyglucose, is used as a tracer to visualize the presence of tumors through positron emission topography (Wechalekar, Sharma et al. 2005).

High levels of glycolytic metabolism do provide a series of advantages to cancer cells.

Firstly, glycolysis generates the necessary precursors for the de novo synthesis of nucleotides and lipids which are required in large quantities during the stages of uncontrolled division and proliferation. (Bui and Thompson 2006). Besides being provided with ample energy and biosynthetic building blocks, cancer cells suppress healthy neighboring cells by consuming the majority of diffusible substrates (Pfeiffer, Schuster et al. 2001). Furthermore, accelerated glycolytic processes causes a drop in the pH of the surrounding tissue which can initiate apoptosis in neighboring cells with a functional p53 (Williams, Collard et al. 1999). Despite the advantages, it must be noted that the Warburg Effect makes the cells dependent on the availability of substrates (Gatenby and Gillies 2004).

PFKFB3 and Cancer

In the absence of stimulator, PFK-1 is inactive in physiological conditions, and tumor cells do contain inhibitory levels of ATP. It is thought that high levels of F-2,6-P2 may contribute to aerobic glycolysis within tumor cells (Hue and Rousseau 1993). Studies of established cancer cell lines have indicated that F-2,6-P2 concentrations are often elevated (Hue and Rousseau 1993). In the case of chronic lymphocytic leukemia cells, the concentration of F-

2,6-P2 is the same as in control lymphocytes, but the sensitivity of PFK-1 for F-2,6-P2 is about one-hundred times greater (Colomer, Vivescorrons et al. 1987).

Due to the reports concerning F-2,6-P2 within cancer biology, an analysis of the bifunctional enzyme’s activity within cancer cells was initiated. These studies showed that bifunctional enzyme taken from cancer cells was seemingly devoid of F-2,6-P2ase activity

(Loiseau, Rider et al. 1988). This would hint to presence of the inducible isoform of the bifunctional enzyme. Indeed, later studies have shown that PFKFB3 mRNA is expressed in breast cancer, leukemia, and adenocarcinoma (Hamilton, Callaghan et al. 1997; Chesney,

Mitchell et al. 1999). PFKFB3 was later found to be neoplastic requirement for transformation by the oncogene, ras (Telang, Yalcin et al. 2006). The suppression of PFKFB3 through the use of silencing siRNA resulted in the reduction of F-2,6-P2 concentrations and glycolytic activity, as well as a reduction in cancer cell viability (Calvo, Bartrons et al. 2006).

The Hypoxia-Inducible Transcription Factor HIF-1

An explanation for the expression of PFKFB3 in cancer cells can be possibly be attributed to the hypoxia-inducible transcription factor HIF-1. HIF-1 is the primary regulator of the hypoxic response in animal cells. Particularly, HIF-1 is responsible for a switch in form of energy metabolism from oxidative phosphorylation to glycolysis. HIF-1 is frequently expressed at high levels in cancer cells, and in turn, induces the expression of genes involved in erythropoiesis, angiogenesis, and anaerobic metabolism (Caro 2001; Maxwell, Pugh et al. 2001;

Brahimi-Horn and Pouyssegur 2006; Semenza 2006). The activation of HIF-1 can be triggered by a series of growth factors and oncogenes (Thornton, Lane et al. 2000; Semenza 2002; Zhou and Brune 2006).

HIF-1 is a hetero-dimeric transcription factor formed by an -subunit, whose expression is dependent of the availability of O2, and a -unit, which is constitutively expressed (Semenza

2002). In the absence of O2, the -subunit is no longer subject to degradation via the ubiquitin-

22 proteasome system (Huang, Arany et al. 1996; Salceda and Caro 1997). The functional heterodimer works by binding hypoxia response elements within the promoters of the targeted genes (Caro 2001; Maxwell, Pugh et al. 2001). In such a manner, HIF-1 suppresses the Krebs cycle and respiration by inducing transcription of pyruvate dehydrogenase kinase 1 (PDK1).

PKD1 reduces the amount of pyruvate entering the Krebs cycle by inactivating pyruvate dehydrogenase though phosphorylation (Kim, Tchernyshyov et al. 2006). This also reduces oxygen consumption in the cell (Papandreou, Cairns et al. 2006). Also included amongst the transcriptional targets of HIF-1 are glucose transporters and glycolytic enzymes including

PFKFB3 (Dang and Semenza 1999; Minchenko, Leshchinsky et al. 2002; Obach, Navarro-

Sabate et al. 2004). Indeed, PFKFB3 is expressed in many cancer cells (Atsumi, Chesney et al.

2002; Calvo, Bartrons et al. 2006; Telang, Yalcin et al. 2006).

PFKFB3 as a Therapeutic Target

Cancer cells are dependent on glycolysis as metabolic source of energy, and it has been observed that cancer cells are sensitive to glucose deprivation. For these reasons, selective inhibition of glycolysis within cancer cells has become a therapeutic target (Shim, Chun et al.

1998). Inhibitors of the glycolytic enzyme, hexokinase, have been successfully shown to suppress tumor growth (Xu, Pelicano et al. 2005; Pelicano, Martin et al. 2006). The studies, which show that siRNA suppression of PFKFB3 results in a reduction of cancer cell viability

(Calvo, Bartrons et al. 2006), has led to attempts to generate selective inhibitors for the inducible isoform of the bifunctional enzyme. One group was able to identify a novel PFKFB3 inhibitor through in silico techniques, 3-(3-pyridinyl)-1-(4-pyridinyl)-2-propen-1-one (3PO). 3PO is capable of reducing cellular concentrations of F-2,6-P2 and glycolytic flux in transformed cells.

3PO is also selectively cytostatic to ras-transformed cells, and has been shown to suppress tumor

23 growth in breast adenocarcinoma, leukemia, and lung adenocarcinoma cells in vivo (Clem,

Telang et al. 2008). A second group has also identified two more PFKFB3 inhibitors, N4A (5, 6,

7, 8-tetrahydroxy-2-(4-hydroxyphenyl) chromen-4-one) and YN1 (7, 8-dihydroxy-3-(4- hydroxyphenyl) chromen-4-one), using in silico techniques(Seo, Kim et al. 2011). These inhibitors as well were capable of reducing cellular F-2,6-P2 concentrations and glycolytic flux in cancer cell lines, which resulted in a suppression of tumor growth and massive cell death.

Additionally, the inhibitors were found within the x-ray crystallographic structure of PFKFB3, which will eventually lead to a foundation for structure assisted optimization and development of novel PFKFB3 inhibitors (Seo, Kim et al. 2011).

CHAPTER 2: MOLECULAR BASIS OF THE FRUCTOSE-2,6-BISPHOSPHATASE REACTION OF PFKFB3: TRANSITION STATE AND THE C-TERMINAL FUNCTION*

Introduction

Fructose-2,6-bisphosphate (F-2,6-P2) is known to be the most potent allosteric activator of the rate-limiting enzyme in glycolysis, 6-phosphofructose-1-kinase (PFK-1), as well as an inhibitor of the gluconeogenic enzyme fructose-1,6-bisphosphatase (F-1,6-P2ase). As such, this small signaling molecule allows for switching between the two pathways of glucose metabolism within the liver, playing a crucial role in maintaining glucose homeostasis in mammalian bodies and controlling the rates of glycolysis in other tissues(Claus, El-Maghrabi et al. 1984). The concentration of F-2,6-P2 is regulated by the bifunctional enzyme, 6-phosphofructo-2- kinase/fructose-2,6-bisphosphatase (PFKFB). The N-terminal half of the enzyme subunit (6- phosphofructo-2-kinase domain) catalyzes the synthesis of F-2,6-P2, using fructose-6-phosphate

(F-6-P) and adenosine 5'-triphosphate (ATP) as substrates (Figure 2.1A). Conversely, the C- terminal half (fructose-2,6-bisphosphatase domain) is responsible for the hydrolytic degradation of F-2,6-P2 into F-6-P and inorganic phosphate (Pi). These two mutually opposing catalytic activities are controlled by different mechanisms such that either activity is predominant in a given physiological condition. Ultimately, the PFKFB control of F-2,6-P2 production/degradation controls the rate of glucose metabolism (Kountz, Freeman et al. 1988).

In order to optimize the regulation of F-2,6-P2 in a tissue-specific manner, there exist four

PFKFB tissue-specific isoforms each encoded by separate genes pfkfb1-4. They are liver,

PFKFB1; heart, PFKFB2; testis, PFKFB4; and inducible forms, PFKFB3 (Algaier and Uyeda

* This chapter first appeared as “Cavalier, M.C.; S.-G. Kim, et al. (2012) Molecular basis of the Fructose-2,6-bisphosphatase reaction of PFKFB3: Transition state and the C-terminal function. Proteins 80, 1143-53”. The “Material” was reproduced with permission of John Wiley & Sons, Inc. ©2011 Wiley Periodicals, Inc.

1988; Sakai, Kato et al. 1996; Heine-Suner, Diaz-Guillen et al. 1998; Manzano, Rosa et al.

1998). These isoforms share highly conserved core catalytic domains (85%) (Figure 2.2), but differ greatly in their kinetic properties and responses to regulatory signals. These differences are mostly due to highly divergent N- and C-terminal regulatory domains(Kurland, Li et al.

1993; Lin, Kurland et al. 1994; Kurland, Chapman et al. 2000); however, a few but significant sequence differences in the catalytic domains that constitute the secondary residue shells surrounding the active sites also contribute to the kinetic differences(El-Maghrabi, Noto et al.

2001). Differences in the structure/function relationships between these isoforms have been

Figure 2.1. Overall folding of PFKFB3 and sites of ligand binding. All figures, unless otherwise noted, were constructed using PyMOL (The PyMOL Molecular Graphics System, Version 1.2r3pre, Schrödinger, LLC.). (A) The overall fold of the monomeric PFKFB3 is represented in a ribbon diagram. The N-terminal kinase domain (6-PF-2-k) is shaded red while the C-terminal bisphosphatase domain (F-2,6-P2ase) is shaded blue. Each active site location can be discerned by the bound ligands. The ADP and F-2,6-P2 are within the kinase active site while F-6-P and a covalently attached phosphate are within the bisphosphatase active site. Molecules of interest are represented in stick models. (B) To compare the organization of the C-termini, the bisphosphatase domains of the inducible (blue) (PDB ID: 2AXN) (Kim, Manes et al. 2006), testes (yellow) (PDB ID: 1BIF) (Hasemann, Istvan et al. 1996), and liver (red) (PDB ID: 1K6M) (Lee, Li et al. 2003) isoforms have been superimposed. The folding patterns are represented by a tracing of the C-alphas.

Figure 2.2. Sequence alignment of PFKFB. Shown are the sequence alignments of the bisphosphatase domain of the four PFKFB isoforms. Residue numbering coincides with the residues of PFKFB3. Residues involved in catalysis, substrate binding, and the initiation of the conformational change are emphasized with boxes. Asterisks mark residues of interest that are conserved in all isoforms except for PFKFB3.

addressed using the available crystal structures of PFKFB1, PFKFB3, and PFKFB4(Hasemann,

Istvan et al. 1996; Lee, Li et al. 2003; Kim, Manes et al. 2006). For a more complete comparison, the structure of PFKFB2 needs to be determined.

Unlike the other isoforms, PFKFB3 is ubiquitously expressed upon the onset of hypoxia(Minchenko, Leshchinsky et al. 2002; Minchenko, Opentanova et al. 2003) and has been shown to be identical to the isoforms isolated from placenta, pancreatic -islet, and brain(Sakakibara, Okudaira et al. 1999; Goren, Manzano et al. 2000; El-Maghrabi, Noto et al.

2001). PFKFB3 has a uniquely large 6-phosphofructo-2-kinase to fructose-2,6-bisphosphatase

27 activity ratio compared to other isoforms. This isoform, which has a native activity ratio of roughly 700 fold kinase-to-phosphatase activity, dramatically increases up to 3000 upon phosphorylation of Ser460 by protein kinase A (PKA) or AMP-dependent protein kinase

(AMPPK)(Okamura and Sakakibara 1998; Marsin, Bertrand et al. 2000; Atsumi, Chesney et al.

2002; Marsin, Bouzin et al. 2002; Manes and El-Maghrabi 2005). Recent studies suggest that this form is a causative molecule of highly elevated levels glycolysis in tumor cells, a phenomenon known as the ‘Warburg effect’(Warburg 1956). Thus, it has also been suggested that PFKFB3 may serve as a promising new target for future cancer chemotherapy(Hirata,

Watanabe et al. 2000; Chesney, Telang et al. 2005). Consequently, an intensive study to investigate the structure/function relationship of PFKFB3 has been performed, and the catalytic mechanism of the kinase reaction for the synthesis of F-2,6-P2 was recently hypothesized from the structural evidence(Kim, Manes et al. 2006; Kim, Cavalier et al. 2007).

The low bisphosphatase activity of PFKFB3, which is lower than that of other isoforms by an order of magnitude(Manes and El-Maghrabi 2005), provides a unique opportunity for the study of the bisphosphatase reaction. In the present study, we determined an improved crystal structure of the PFKFB3 phospho-enzyme intermediate state (PFKFB3-P•F-6-P) trapped during the bisphosphatase reaction; PFKFB3 complexed with aluminum tetrafluoride (AlF4) within the bisphosphatase catalytic pocket (PFKFB3•AlF4), which mimics the transition state of a phosphoryl transfer; and PFKFB3 complexed with inorganic pyrophosphate (PFKFB3•PPi) in which a novel conformation of C-terminus residues 440-446 was observed. From these structures, we were able to deduce all the structures necessary for understanding of this reaction, namely, the Michaelis complex state and the transition states. This study provided a new insight into the catalytic mechanism of this histidine phosphatase and the results are discussed here.

Results

Overall Structure

In this study, three models (PFKFB3-P•F-6-P, PFKFB3•AlF4, and PFKFB3•PPi) which in concert detail the enzyme state through substrate binding, catalysis, and product release during the bisphosphatase reaction (Figure 2.3) are being introduced. The global conformation of these models including crystal contacts, the dimeric interface, and domain/domain interactions are not significantly affected by the liganding conditions varied in this experiment. As detailed in previous structural studies, a subunit of the active dimer of PFKFB3 is comprised of a single polypeptide encompassing two catalytic domains. (Figure 2.1A) In each complex, N-terminal residues 4 through 15 form a β-hairpin structure to make contacts with bisphosphatase domain at the area where the residues involved in binding of both product and substrate are concentrated.

In this way, the self-regulatory effect of the N-terminus on F-2,6-P2ase activity(Kim, Manes et al. 2006) is not affected by experimentally varied liganding. Unlike the N-terminus, the C- terminus residues (460-520) remain disordered and have not yet been elucidated. The only apparent conformational change observed occurs at C-terminus residues 440-446 and is explained in detail in later sections of this article.

PFKFB3-P•F-6-P Structure

The electron densities of the crystals soaked with F-2,6-P2 show F-2,6-P2 to be dephosphorylated to F-6-P within the catalytic pocket of the bisphosphatase domain. The phosphate transferred from O2 of F-2,6-P2 was covalently bound to NE2 of His253 with a distance of 1.7Å in a covalent phospho-enzyme intermediate state (PFKFB3-P•F-6-P). The distance between the His-P phosphorous and O2 of F-6-P, the leaving group, is 3.4Å. The oxygens of the His-P phosphate make numerous interactions with the protein residues such that the E-P state is stabilized. The residues involved in the interactions with the E-P phosphate are 29

Figure 2.3. The minimal enzymatic reaction. The various enzyme states of PFKFB3 through a minimal enzymatic reaction are shown: free enzyme (E), the Michaelis complex (E•F-2,6-P2), phospho-enzyme intermediate (E-P•F-6-P), and the states of product release (E-PE•Pi).

Arg252, His387, and Asn259. The details of these interactions are described in following sections, and they are summarized in Table 2.1. The improved PFKFB3-P•F-6-P model described here is an extension of resolution and confirms the orientation previously described by

PDB ID:2I1V(Kim, Cavalier et al. 2007). A similar model of PFKFB in the E-P•F-6-P intermediate state has been previously described using the rat liver isoform (RL2K) of

PFKFB(Lee, Olson et al. 1997). Unfortunately, conclusions regarding the human PFKFB3 should be drawn cautiously from the RL2K model, as this was a truncated form, which lacked the kinase domain and thirty residues of the C-terminus. As a result, the bound F-6-P was found near the interface of two protein molecules within the crystal lattice, and the water- bridgeinteractions between the analogous Thr440 and F-6-P, whose importance is described in later sections, was lost. These circumstances resulted in a notably different position of F-6-

P(Lee, Olson et al. 1997; Yuen, Mizuguchi et al. 1999).

PFKFB3AlF4 Structure

A crystal soaked with NaF and AlCl3 shows an electron density representing a square planar AlF4 within the catalytic pocket of the PFKFB3 bisphosphatase domain (Figure 2.4A).

The metal makes two collinear axial bonds to NE2 of His253 and to the oxygen of a water molecule, each with the distance of 2.1Å and 1.9Å, respectively. As shown in Figure 2.4A, F3 interacts with His387, F4 with Asn259 and Arg252, and F2 with Arg252. F1 of AlF4 makes a

Table 2.1. Notable interactions between the bisphosphatase active site ligands and the protein.

Protein Atoms His-P AlF4 PPi F6P Arg252 NH1 O1 (3.0) F4 (2.8) O4 (3.0) O3 (3.2) F2 (3.2) O5 (3.3) NE O3 (2.9) F2 (2.7) O5 (3.0) His253 NE2 P (1.7) * Al (2.1) P2 (3.5) Asn256 ND2 F1 (3.5) O3 (3.0) Asn259 OD1 O1 (3.1) F4 (3.0) O4 (3.0) Glu322 OE2 O6 (2.6) O2 (2.5) His387 ND1 O2 (2.8) F3 (2.6) O (3.3) Tyr333 OH O3P (2.6) Arg347 NH1 O3P (2.7) NH2 O1P (2.9) Tyr362 OH O1P (2.5) Arg392 NH1 O2P (2.6) Values in parentheses are distances given in angstroms. * Indicates a covalent bond.

unique short-range interaction with Asn256. Geometric details of the interactions are provided in Table 2.1.

Consequently, the atoms of bound AlF4, together with the apical water and NE2 of

His253 constitute, a tetragonal bipyramidal structure, which seems to mimic a trigonal bipyramidal phosphorane structure that is transiently formed during the phosphoryl transfer reaction. The apical water molecule also makes a short-range, 2.8Å, interaction with Glu322.

This finding strongly suggests the function of Glu322 is to neutralize the leaving group and to activate the attacking water molecule.

The PFKFB3•AlF4 structure is the first PFKFB3 model described in which the product of the phosphatase domain, F-6-P, is absent. It is also the first structure in which the invariant region (residues 440-446), whose amino acid sequence and conformation is strongly conserved

Figure 2.4. Aluminum tetrafluoride within the active site. (A) A stereo view within the bisphosphatase active site of the PFKFB3•AlF4 model is shown. The mesh represents the |Fo| - |Fc| electron density omit map of aluminum tetrafluoride and a bound water molecule within the bisphosphatase active site. The map is contoured at 3.0levels. Arg252, Asn259, and His387 interact with the peripheral fluorines of aluminum tetrafluoride. Glu322 interacts with a water at the apex of the bipyramidal structure. The aluminum atom is situated 1.9Å from the water and 2.1Å from His253. The dotted lines mark significant interactions. Molecules/residues of interest are represented in stick models. (B) Shown is a ribbon representation in stereo view of the overlapping bisphosphatase active sites of the PFKFB3-P•F-6-P(green) and PFKFB3•AlF4 (yellow) models. C-terminal residues 440-446 of the PFKFB3•AlF4 model can no longer be tracked. Spheres represent the positions of the C-alphas of the missing residues as projected from the PFKFB3-P•F-6-P model. among all four PFKFB isoforms, is disordered (Figure 2.4B). Details regarding this particular conformational change will be discussed in the following sections.

PFKFB3•PPi Structure

The crystal soaked with pyrophosphate and F-6-P showed electron density representing pyrophosphate (PPi) within the bisphosphatase site (Figure 2.5A), while the kinase site was occupied by two pyrophosphate molecules (data not shown). Within the bisphosphatase domain, the second phosphorous of the pyrophosphate and its conjugate oxygens have a lower occupancy. The high occupancy phosphorus atom is 3.5Å from His253. Although it was included in the soaking solution, F-6-P is absent from the bisphosphatase active site, suggesting that the E•Pi•F-6-P complex is not favored. This model can be taken as a representative of the

E•Pi state.

Residues 440-446, which were interpreted as a random coil in all models before the

PFKFB3•AlF4 structure, in which it was disordered (Figure 2.5C), can now be seen as a weak, continuous density which was interpreted as an -helix extending outward from the active site

(Figure 2.5C).

Comparison of Bisphosphatase Active Sites

Three models are described above (PFKFB3-P•F-6-P, PFKFB3•AlF4, and PFKFB3•PPi).

Within each of these three models, a representation of the transferred 2-phosphate (His-P, AlF4, and PPi, respectively), each at a various stage of the catalytic reaction (E-P•F-6-P, E-P-F-6-P/E-

P-H2O transition states, and E•Pi, respectively), can be extracted. Within each of these representations, the represented oxygens of the transferred 2-phosphate maintain interactions with His387, Asn259, and Arg252 (Table 2.1). Therefore, these residue side-chains must remain in contact with the 2-phosphate oxygens throughout the reaction.

Glu322 makes interactions with the O2 of F-6-P in the PFKFB3-P•F-6-P model, with the apical water of the bipyramid within the PFKFB3•AlF4 model, and with the O1 of high

Figure 2.5. Pyrophosphate within the active site. (A) A stereo view within the bisphosphatase active site of the PFKFB3•PPi model is shown. The mesh represents the |Fo| -|Fc| electron density omit map of PPi within the bisphosphatase active site. The map is contoured at 3.0levels. Arg252, Asn259, and His387 interact with the pyrophosphate oxygens that are assumed to be at the positions of the 2-phosphate oxygens of F-2,6-P2. The phosphorous is situated 3.5Å from His235. The dotted lines mark significant interactions. Molecules/residues of interest are represented in stick models. (B) Ribbon representations in stereo of the overlapping models of PFKFB3-P•F-6-P (green) and PFKFB3•PPi (magenta) are shown. Molecules/residues of interest are represented in stick models. Upon the absence of F-6-P, two waters which bridge interactions with the C-terminal segment covering the active site are lost. The C-terminal residues 440-446 are no longer seen as a random coil extending over the bisphosphatase active site, but as a helix extending outwards away from the active site. After the loss of the two water bridged interactions, Thr440 rotates out of the active site. Simultaneously, His441 replaces Arg442 in a -cation interaction with Tyr333. Arg442 can then commence the coiling of the observed helix. (C) Shown are the C-terminal residues of the PFKFB3•PPi model. Residues 425-436 are represented as a ribbon while residues 437-446 are represented with sticks. The mesh represents the |Fo| -|Fc| electron density omit map of C- terminal residues 437-446 of the bisphosphatase active site. The map is contoured at 2.0levels.

35 occupancy phosphate of PPi within the PFKFB3•PPi model (Table 2.1). The significance of this will be explained in later sections.

The O1 oxygen of the high occupancy phosphate of PPi within the PFKFB3•PPi model is at the approximately same position as the O2 of F-6-P within the PFKFB3-P•F-6-P model. The summation of the atoms within these two molecules yields a representation of the bisphosphatase substrate F-2,6-P2 (Figure 2.6A), allowing a hypothetical model of the Michaelis complex

(Figure 2.6B).

Because of the absence of F-6-P within the PFKFB3•PPi model, two waters, which bridge extensive interactions between F-6-P and the C-terminus in the PFKFB3-P•F-6-P model, are lost. The C of Thr440 moves 2.5Å out of the active site, and the side-chain rotates 113-132° outward from the active site. Simultaneously, His441 acts to replace Arg442 in a -Cation interaction with Tyr333. The displacement of Arg442 allows coiling of the newly observed - helix (Figure 2.5B).

Degradation of Fructose-2,6-Bisphosphate

Based on the three structures, a molecular model of the F-2,6-P2ase reaction of the human inducible form of PFKFB is suggested. This mechanism is in direct agreement with the mechanism suggested for the human testes form of PFKFB(Yuen, Mizuguchi et al. 1999). The models introduced in this work, in particular PFKFB3AlF4, increase confidence in assumptions concerning the mechanism of PFKFB3 by offering structural details that were not previously available in any isoform.

Binding of Substrate (E  E•F-2,6-P2). In Figure 2.6B, the F-2,6-P2 representation summed from the high occupancy phosphate of the PFKFB3•PPi model and the F-6-P of the

PFKFB3-P•F-6-P model confirms that the 6-phosphate moiety is recognized by Arg347, Lys351,

Figure 2.6. A hypothetical representation of the Michaelis complex (E•F-2,6-P2). Here is the Michaelis complex as constructed using ligands from the PFKFB3-P•F-6-P and PFKFB3•PPi models. Molecules/residues of interest are represented in stick models. (A) A stereo view of the overlapping bisphosphatase active sites of both the PFKFB3-P•F-6-P(green) and PFKFB3•PPi(magenta) models is shown. The pyrophosphate of the PFKFB3•PPi model and the F-6-P of the PFKFB3-P•F-6-P model have been included but not the covalently bound phosphate. The lower occupancy phosphoryl group of PPi has been grayed. The remaining ligand atoms can be used to construct a continuous representation of bound F-2,6-P2. (B) A complete representation of the Michaelis complex projection is shown in pink. The 2-phosphate moiety is recognized by residues Arg252, Asn259, and His387. The 6-phosphate moiety is recognized by residues Arg347, Lys351, and Tyr362. (C) The ligands from all three models are displayed. The lower occupancy phosphoryl group of PPi has been grayed. The direct inversion of oxygen stereochemistry, through the E•F-2,6-P2 to E-P states with the transition state represented by the equatorial plane formed by the peripheral fluorines of AlF4, can be tracked. By tracking through all three representations of the 2-phosphate phosphorous (His-P, AlF4, PPi), one would create an approximately straight line punctuated at the ends by the NE2 of His253 and the O2 of F-6-P.

37 and Tyr362 and that the 2-phosphate moiety is recognized by residues His387, Asn259, and

Arg252. The C-terminal residues 440-446 are stabilized as a random coil through water- mediated interactions with F-2,6-P2.

Formation of the Phospho-Enzyme Intermediate (E•F-2,6-P2  E-P•F-6-P). In Figure

2.6C, the three newly introduced models allow us to observe three positions of the 2-phosphate during the phosphoryl transfer; PPi, AlF4, and His-P. By tracking through all three representations of the phosphorous of 2-phosphate (His-P, AlF4, PPi), a nearly straight line punctuated at the ends by the NE2 of His253 and the O2 of F-6-P can be seen, suggesting an “in- line” transfer mechanism(Kountz, Freeman et al. 1988). This supports the direct inversion of oxygen stereochemistry through the PFKFB3-P•F-6-P to PFKFB3•PPi models with the transition state represented by the equatorial plane formed by the peripheral fluorines of AlF4. An equatorial plane created by the migrating peripheral oxygens of phosphate was hypothesized

(Yuen, Mizuguchi et al. 1999), but never before has it been so clearly depicted. Note that the arrangement of AlF4, bound by PFKFB3, cannot exactly mimic the trigonal bipyramidal transition state of the phosphoryl-transfer reaction but is considered an adequate approximation in phosphoryl-transfer mechanisms(Coleman, Berghuis et al. 1994; Fisher, Smith et al. 1995;

Rittinger, Walker et al. 1997; Olesen, Sorensen et al. 2004; Golicnik 2010).

Upon binding of F-2,6-P2, the NE2 of His253 is in position for a nucleophilic attack on the 2- phosphate. This attack is aided by the interactions of 2-phosphate with Arg252, His387, and

Asn259. The resulting interactions would stabilize the transition state by delocalizing the electrons in the 2-phosphate towards the side-chains of residues Arg252, His387, and Asn259 making the phosphorous of the 2-phosphate more electrophilic. This would enhance the nucleophilic attack of the lone pair of electrons of the NE2 of His253. The removal of 2-

38 phosphate from F-2,6-P2 would leave a highly basic anion that would need to be neutralized to prevent reformation of the reaction substrates. The general proximity of Glu322 suggests that it is positioned to function as the general acid and donates a proton to the leaving O2 after His-P formation. This is consistent with kinetic studies of the conserved Glu327 within the rat liver isoform, which suggests that Glu327 acts as both a general acid during the E-P formation and a general base during E-P breakdown(Kountz, Freeman et al. 1988; Sakurai, Hasemann et al.

2000). The reaction mechanism resulting in the E-P•F-6-P intermediate is summarized in Figure

2.7.

Release of F-6-P and Degradation of the Phospho-Histidine Intermediate (E-P•F-6-P 

E-P  E•Pi). Upon the departure of the F-6-P and the two waters which bridge interactions between F-6-P and residues 440-446, this segment of the C-terminus is no longer stabilized as a random coil. A molecule of water would then move in to the apex of a tetragonal bipyramidal configuration and the mechanism by which E-P would breakdown is presumably the reverse of its formation. Glu322 would withdraw a hydrogen from the water and the resulting strong anion would attack the electrophilic phosphate while the transition state is stabilized by residues

Arg252, His387, and Asn259 (Figure 2.7). The importance of the water found at the apex of the bipyramid now becomes immediately apparent. The activation of water by Glu322 has been hypothesized (Kountz, Freeman et al. 1988; Yuen, Mizuguchi et al. 1999; Sakurai, Hasemann et al. 2000), but the placement of a water molecule in a position of attack has never before been depicted. The track of the peripheral oxygens through the equatorial transition state during the formation of free phosphate is also represented by the PFKFB3•AlF4 model. In actuality, because of the absence of F-6-P in the PFKFB3•AlF4 model, the track of the peripheral oxygens during the phospho-histidine breakdown is more directly portrayed than the formation of the

Figure 2.7. The complete mechanism of phospho-histidine formation/degradation. This schematic was prepared with ChemBioDraw (ChemBioDraw Ultra 10.0 CambridgeSoft, 100 CambridgePark Drive, Cambridge, MA 02140). Brackets indicate a transition state in which a direct inversion of phosphate stereochemistry occurs. Upon binding of F-2,6-P2, His253 is in position for a nucleophilic attack on the 2-phosphate. Interactions with Arg252, His387, and Asn259 stabilize the transition state by delocalizing the electrons towards the residues making the P of 2-phosphate more electrophilic. This enhances the nucleophilic attack of the lone pair of electrons of the NE2 of His253. Glu322 acts as a general acid and donates a proton to neutralize the highly basic anion on the first product. Glu322 would then withdraw a hydrogen from a water, and this strong anion would attack the electrophilic phosphate while the transition state is stabilized by residues Arg-252, His-387, and Asn-259 creating the second product

phospho-histidine. Upon the formation of the free phosphate, residues 440-446 shift away from the active site and appear to coil into an -helix.

Discussion

The three models introduced in this article provide a clearer understanding of the bisphosphatase reaction of the inducible form of the bifunctional enzyme by both supporting previous assumptions derived from other isoforms, and distinguishing unique characteristics of

40 this particular polypeptide. The PFKFB3-P•F-6-P model allows confirmation of the positioning of the fructose and 6-phosphate atoms as well as the residues responsible for their binding, whereas the PFKFB3•PPi does the same for the atoms of the 2-phosphate. The PFKFB3•AlF4 model directly illustrates the stereochemistry of the bisphosphatase reaction for the first time amongst all isoforms.

It has been suggested that the reduced bisphosphatase activity of PFKFB3 is solely due to the presence of a serine at residue 302 instead of an arginine as conserved in the other isoforms

(Figure 2.2). This residue is said to interact with the 2-phosphate and further stabilizes the transitions state, if conserved(Hasemann, Istvan et al. 1996; Lee, Li et al. 2003; Kim, Manes et al. 2006). Studies have reported that the restoration of this serine to arginine rescues the ability to generate and degrade the His-P intermediate(Manes and El-Maghrabi 2005). Strangely, however, despite not a having arginine at position 302, PFKFB3 has the ability to form the phospho-histidine intermediate but merely lacks the ability to degrade it. This article provides structural evidence of a conformational change in C-terminal residues

440-446 and suggests that the reason for the depressed bisphosphatase activity may be more complex than simply a single mutated residue. The absence of F-6-P from the active site causes rearrangement of the C-terminus residues 440-446. Perhaps, this is suggestive that bisphosphatase activity is further regulated through the control of ligand binding and product release, or perhaps the degradation of the active site after the release of F-6-P.

It should be noted that the pivotal residues of this conformational change (Thr440,

His441, Arg442) are absolutely conserved across all known mammalian isoforms of fructose-

2,6-bisphosphatase. In addition, previous works with the rat liver isoform, which was truncated just before the Thr-His-Arg segment, resulted in a nine-fold increase in kcat, an eleven-fold

41 increase in Km, and relief of pH dependent regulation of bisphosphatase activity(Lin, Kurland et al. 1994).

No similar conformation change in residues 440-446 has been seen in any other isoforms

(Figure 2.1B), yet additional studies are needed to conclusively ensure that this phenomenon is unquestionably unique to PFKFB3. However, from our research, we are confident that the - cation interaction between Arg442 and Tyr333 is unique to PFKFB3 among all currently known isoforms. In both PFKFB1 and PFKFB4 models, this conserved arginine forms a salt bridge with an aspartic acid, namely Asp353 and Asp351, respectively. Arg442 does not interact with a glutamic acid, Glu348, at this same position within PFKFB3(Figure 2.2).

Materials and Methods

Preparation and crystallization of PFKFB3

Protein preparation and crystallization was similar to methods previously described(Kim,

Manes et al. 2006; Kim, Cavalier et al. 2007). The His6-tagged human inducible isoform of the bifunctional enzyme, PFKFB3, was expressed in Escherichia coli BL21 (DE3) pLysS and was purified by Ni-NTA affinity columns. The protein sample was further purified using Q- sepharose anion-exchange chromatography. This purified protein was kept at a concentration of

-1 8 mg ml in pH 8.0, 20 mM Tris-HCl, 10 mM NaxPi, 5 mM -mercaptoethanol, and 5% glycerol. Crystal were grown using sitting drop vapor diffusion with a 1:1 (v/v) mixture of protein with a mother liquor of 100 mM Tris-HCl, pH 7.5, 20-25% ethylene glycol, 200-400mM

Tartaric Acid, 5% glycerol, and 12% polyethylene glycol 4000. Crystals in a size of 0.2 X 0.2 X

0.05 mm were grown in two to three weeks.

The crystals were soaked with cryoprotectant solutions for 0.5 to 2 hours to allow binding of the desired ligands. Cryoprotectant solutions consisted of 20 mM Tris-HCl pH 7.5 buffer and

12% polyethylene glycol 4000, and were enriched with 35% ethylene glycol and the targeted ligands. The PFKFB3-P•F-6-P crystal was soaked with 3mM ADP 0.5mM F-2,6-P2. The

PFKFB3•AlF4 crystal was soaked with 3mM ADP, 0.5mM F-6-P, 2mM AlCl3, and 8mM NaF.

The PFKFB3•PPi crystal was soaked with 3mM PPi. The soaked crystals were flash cooled at

100K and the temperature was maintained during data collection using an Oxford cryo device.

Data Collection and Processing

Diffraction data for the PFKFB3•AlF4 and the PFKFB3-P•F-6-P crystals was collected at

The Gulf Coast Consortium Protein Crystallography Beamline (PX1) in The Center for

Advanced Microstructures and Devices (CAMD), Louisiana State University, Baton Rouge, LA using a x-ray source wavelength of 1.3808Å the data was recorded on a Mar 165mm CCD detector and was integrated, merged, and scaled using HKL2000(Otwinowski and Minor 1997).

Diffraction data for the PFKFB3•PPi crystal was collected at The Northeastern

Collaborative Access Team (NE-CAT) beamline at the Advanced Photon Source (Argonne

National Laboratory) using an x-ray source wavelength of 0.9792Å. The data was recorded on an ADSC Q315 (315mm X 315mm) detector and was integrated, merged, and scaled using

HKL2000(Otwinowski and Minor 1997). Statistics of the diffraction data and structure refinement are summarized in Table 2.2. All crystals belonged to the P6522 space group, and the cell dimensions were similar.

Structure Determination and Refinement

The reduced data was formatted for the program suite of CCP4(Bailey 1994; Potterton,

Briggs et al. 2003)and 5% of the data was marked for free R-factor measurements in subsequent structure refinements. Initial models of all complexes were determined using REFMAC(Vagin,

Steiner et al. 2004) within the CCP4 suite with the PFKFB3•ADP•EDTA complex (PDB ID:

Table 2.2. Statistics of reflection data and structure refinements (PFKFB3). Liganding PFKFB3•AlF4 PFKFB3-P•F-6-P PFKFB3•PPi Space group P6522 P6522 P6522 Unit cell dimension (Å) 102.71 X 102.71 X 257.04 102.15 X 102.15 X 259.44 102.98 X 102.98 X 258.13 Wilson Plot B Value (Å2) 45.7 55.6 41.2 Resolution range (Å) 30.0 - 2.25 30.0 - 2.45 35-2.30 Reflections Observed 351,069 434,030 927,757 Unique Reflections 38,530 30,381 36,888 Reflections Rfree set 1,951 5,839 1,975 Completeness (%) 99.5 (98.5) 99.6 (98.7) 100 (99.9) Redundancy 4.7 (4.6) 5.8 (5.7) 7.6 (7.8) |/ Σ Ih,j), where h=set of Miller indices, j=set of observations of reflection h, and =the mean intensity. R.M.S.D. values are deviation from ideal values. Rcrys= Σh||Fo,h| – |Fc,h||/ Σh|Fo,h|. Rfree was calculated using 5% of the complete data set excluded from refinement. The numbers in parentheses represent values from the highest resolution shell (2.33-2.25 Å for E•AlF4 complex, 2.54-2.45 Å for the E-P•F-6-P complex, and 2.34-2.30 Å for the E•PPi complex).

2AXN(Kim, Manes et al. 2006)) as the starting molecule after removal of the ligand and solvent molecules. The initial model went through many cycles of manual model rebuilding and validation using the program COOT(Emsley and Cowtan 2004). Binding of the ligands was confirmed, referring to the |Fo| -|Fc| omit maps that were generated, when Rcrys/Rfree reached

0.25/0.28 or below and the ligands were incorporated into the complex models.

As summarized in Table 2.2, the PFKFB3-P•F-6-P complex has Rfree/Rcrys of 0.247/0.193 using a total of 3,849 scatterers, including solvent molecules, against all 30,381 available reflections in the resolution range of 30.0–2.45Å. The structure contains a total of 441 amino acid residues of the full-length protein of 520 residues. As in the PFKFB3•ADP•EDTA complex, the C-terminus (residues 446–520) is mostly disordered.

The final structure of the PFKFB3•AlF4 complex has Rfree/Rcrys of 0.242/0.200 using a total of 3,837 scatterers, including solvent molecules, against all 38,530 available reflections in the resolution range of 30.0–2.25Å. The structure contains a total of 431 amino acid residues of the full length protein of 520 residues.

The final structure of the PFKFB3•PPi complex has Rfree/Rcrys of 0.236/0.184 using a total of 3,959 scatterers, including solvent molecules, against all 36,888 available reflections in the resolution range of 35.0–2.30Å. The structure contains a total of 439 amino acid residues of the full-length protein, which consists of 520 residues.

The atomic coordinates and structure factors of the PFKFB3-P•F-6-P complex (PDB ID:

3QPV), the PFKFB3•AlF4 complex (PDB ID: 3QPW), and the PFKFB3•PPi complex (PDB ID:

3QPU) have been deposited in the RCSB Protein Data Bank.

CHAPTER 3: REVIEW OF LITERATURE, VLDE

Diversification of Natural Products

Natural Products Offer Unique Scaffolding

Natural products continue to play a large role in the drug discovery and development processes. According to a survey of clinically available therapeutic agents, a very large percentage of drugs are in fact natural products or at least have clearly discernible connection to natural products (Danishefsky 2010; Newman and Cragg 2012). Although the prevalence of natural compounds as drugs is not entirely understood, certain advantages of natural occurring materials can be inferred. Firstly, natural products are biosynthesized by their hosts to interact with proteins, such as enzymes or receptors. These types of biomolecules are generally the targets of therapeutic agents. Additionally, natural products by definition are housed in a living host. Drug candidates often fail due to the incompatibility with biological hosts, thus the crude biocompatibility of natural products can be quite desirable (Danishefsky 2010). It should also be noted that the design of potential therapeutics by chemical synthesis is usually weighted by

“drug-likeness”. This tends to lead researchers in a direction that is non-analogous to the composition of natural products since the chemical compositions of natural drugs are often counter-intuitive. Correspondingly, Lipinski's “Rules of Five” often do not apply to natural products when considering “druggable chemical entities” (Lipinski 2004; Keller, Pichota et al.

2006; Macarron 2006). Therefore, natural products often offer a complex and non-intuitive scaffolding ideal for drug development.

Unfortunately, biologically active natural products often possess a valuable, though not necessarily optimized, pharmacophoric space (Danishefsky 2010). Also, some natural product substructures are effectively inaccessible to chemical modification and the total chemical

46 synthesis of a natural product can be a daunting task. Thusly, perturbations to the chemical structure of natural products through other means must be investigated. Investigations into the biosynthetic pathways of these natural products may provide the necessary opportunities to produce novel analogs of natural products via biosynthetic approaches. Already, studies of the biosynthetic pathways of natural products have led to the diversification of the antibacterial erythromycin, the anthelmintic avermectins, and the antitumor indolocarbazoles (Figure 3.1).

The diversifications of these scaffolds were achieved through the genetic manipulation of biosynthetic gene clusters along with the use of alternative biosynthetic precursors, and recombinant proteins for chemoenzymatic synthesis (McDaniel, Thamchaipenet et al. 1999;

Cropp, Wilson et al. 2000; Sanchez, Mendez et al. 2006; Mahmud, Flatt et al. 2007). These studies provide examples of how the understanding of a biochemical pathway can lead to the diversification of a group of biologically important compounds.

Diversification of the Erythromycin Scaffold by Genetic Manipulation

The polyketide erythromycin and its semisynthetic derivatives are the third most widely used class of antibiotics in the world (Figure 3.1) (McDaniel, Thamchaipenet et al. 1999).

Erythromycin has been in clinical use for over 50 years and was first isolated by McGuire et al. in 1952 (Mcguire, Bunch et al. 1952; Pal 2006). The specific antibacterial action of erythromycin is attributed to its ability to obstruct protein synthesis by inhibiting ribosomal activity (Vester and Douthwaite 2001). Due to the complexity of its chemical structure, the total chemical synthesis of this important antibiotic by R. B. Woodward and his colleagues took more than a decade to elucidate (Woodward, Logusch et al. 1981; Pal 2006). R. B. Woodward was followed by a series of medicinal chemists who prepared analogs leading to a second generation of macrolide antibiotics such as clarithromycin, azithromycin, and others (Chu 1995).

Figure 3.1. Differentiated Scaffolds. Shown are the chemical structures of natural products whose scaffolds have been differentiated by methods resulting from the study of biosynthetic pathways.

Eventually, these efforts exhausted the chemical modifications available at the existing functional groups of the macrolide ring, but most of the ring was effectively inaccessible to chemical modification (McDaniel, Thamchaipenet et al. 1999). Upon the discovery of the programmed nature of modular polyketide biosynthesis (Cortes, Haydock et al. 1990; Donadio,

Staver et al. 1991), diversification of polyketides could then be accomplished using genetic engineering strategies (Mcdaniel, Ebertkhosla et al. 1995; Katz 1997).

The modular polyketide synthases are encoded by a cluster of contiguous genes and are an organization of catalytic domains that both bind and modify the polyketide backbone. Each module consists of a ketosynthase, an acyltransferase and an acyl carrier protein (O'Hagan 1991;

McDaniel, Thamchaipenet et al. 1999). The acyl carrier protein catalyzes the extension of the growing polypeptide chain while the choice of 2-carbon extender in the form of a CoA activated small organic acid thioester is selected by the specificity of an acyltransferase (Oliynyk, Brown et al. 1996; Liu, Thamchaipenet et al. 1997; Ruan, Pereda et al. 1997). The final state of each 2- carbon extender is determined by the additional presence of varying catalytic domains in the module such as ketoreductases, dehydratases, and enoyl reductases. In summary, the composition of each module encodes the structure of each 2-carbon unit while the order of modules forms a template for the polyketide product (McDaniel, Thamchaipenet et al. 1999).

The discovery that the structural diversity of polyketides is governed by a sequence of catalytic modules made it feasible to contemplate modifications to the chemically intractable sites of such molecules by genetic engineering. Using such techniques, McDaniel,

Thamchaipenet et al. created a library of structurally diverse polyketides. Using the polyketide synthase responsible for the production of the macrolide ring of erythromycin as a starting point, combinatorial biosynthetic strategies were utilized to incorporate components from different

49 polyketide pathways. The diversity of the generated library would be prohibitively impractical to attempt by means of chemical synthesis (McDaniel, Thamchaipenet et al. 1999).

Diversification of the Avermectin Scaffold by Alternative Precursors

The avermectins are a group of anthelmintic macrolides produced by Streptomyces avermitilis (Figure 3.1) (Campbell and Benz 1984). Like other polyketides, the avermectins are synthesized by a modular polyketide synthase using decarboxylative condensations to grow a polyketide chain from a starter unit with catalysis, extender unit selection, and extender stereochemistry governed by the catalytic domains comprising each module (Katz 1997).

Previous studies aimed at diversification of the polyketide scaffold, such as that of erythromycin

(McDaniel, Thamchaipenet et al. 1999), focused primarily on combinations of different polyketide synthase components to produce perturbations in extender chemical structure (Stassi,

Kakavas et al. 1998; McDaniel, Thamchaipenet et al. 1999). Dutton, Gibson, et al. extended the structural diversity of polyketide scaffolds through the incorporation of different polyketide biosynthetic starter units (Dutton, Gibson et al. 1991).

The extension of the avermectin backbone initiates from a starter unit comprised of either

S(+)--methylbutyryl-CoA or isobutyryl-CoA. One step in the formation of the acyl-CoA form of the -branched-chain fatty acid starter units is a branched-chain -keto acid dehydrogenase reaction (Dutton, Gibson et al. 1991; Denoya, Fedechko et al. 1995). A S. avermitilis mutant, which lacks a functional branched-chain -keto acid dehydrogenase, was isolated (Hafner,

Holley et al. 1991). This mutant could only synthesize natural avermectins when the - branched-chain fatty acids were added to the fermentation medium. However, the addition of alternative fatty acids resulted in the formation of a series of novel avermectins (Dutton, Gibson

50 et al. 1991). The efforts necessary to reproduce this series of avermectins through chemical synthesis would be staggering.

Diversification of the Antitumor Indolocarbazoles by Recombinant Enzymes

The indolocarbazoles are an important class of natural products isolated from actinomycetes, cyanobacteria, slime molds, and marine invertebrates (Gribble 1993; Knolker and

Reddy 2002). The indolocarbazoles exhibit antitumor and neuroprotective activities through the inhibition of protein kinases and DNA topoisomerases, and through direct DNA intercalation

(Akinaga, Sugiyama et al. 2000; Fischer, Rodriguez et al. 2002; Prudhomme 2003). The indolocarbazoles rebeccamycin and staurosporine are derivatives of the indolo[2,3-a]pyrrolo[3,4- c]carbazole ring system to which a sugar residue is attached (Figure 3.1). These indolocarbazoles differ in the attachment of the carbohydrate and in the composition of the pyrrole moiety, and these differences are essential for target selectivity (Sanchez, Mendez et al. 2006). Upon the discovery of the rebeccamycin (reb) and staurosporine (sta) biosynthetic gene clusters (Onaka,

Taniguchi et al. 2002; Onaka, Taniguchi et al. 2003; Sanchez, Mendez et al. 2006), the stage was set to elucidate the steps in the biosynthesis of indolocarbazoles and for the generation of novel indolocarbazole analogs by genetic engineering.

Sanchez, Mendez et al. dissected and reconstituted the entire biosynthetic pathway for rebeccamycin in a Staphylococcus epidermidis host (Sanchez, Zhu et al. 2005). By rationally expressing different combinations of reb genes, numerous rebeccamycin derivatives were produced within the genetically engineered host. Later, sta genes responsible for the differentiation of the indolocarbazole’s carbohydrate components were also introduced.

Selective expression of sta genes lead to further derivatization of the indolocarbazole scaffold including alternative glycosylation by a number of deoxysugars (Sanchez, Mendez et al. 2006).

Because of the potential therapeutic applications of indolocarbazole family of natural products, the demonstrated ability to synthesize indolocarbazole derivatives through biological processes within a convenient bacterial host can prove to be an invaluable tool.

The Aminocyclitol Class of Natural Products

The Variety of Aminocyclitols

Another family of natural products whose biosynthesis is being studied in hopes that such knowledge could lead to the development of clinically important compounds is the aminocyclitols. The aminocyclitols are a class of microbially generated natural products with many clinical and agricultural applications (Mahmud 2003; Davies 2006). Included amongst the aminocyclitols are the aminoglycoside antibiotics, the C7N-aminocyclitols, and the five- membered ring aminocyclitols or cyclopentitols. The aminoglycoside antibiotics were among the first clinical antibiotics and remain an important group of drugs for the treatment of infections (Prasad, Verma et al. 2006; Yamamura, Kawada et al. 2008). The aminoglycoside antibiotics work through the inhibition of the bacterial ribosome and thusly the disruption of protein synthesis. The C7N-aminocyclitol and the cyclopentitol groups of natural products are structurally more diverse and display a variety of biological activities. Amongst the C7N- aminocyclitol family is the antifungal, antibiotic validamycin A. The aminocyclitols are derived biosynthetically from simple sugar units which are in most cases formed by a family of enzymes known as sugar phosphate cyclases (SPCs) (Llewellyn and Spencer 2006; Mahmud, Flatt et al.

2007; Wu, Flatt et al. 2007). The aminoglycoside antibiotic precursors are produced by the 1- myo-inositol 1-phosphate synthase and the 2-deoxy-scyllo-inosose synthases while the 2-epi-5- epi-valiolone synthases are involved in the biosynthesis of the C7N-aminocyclitol natural products. Uniquely, the five-membered ring aminocyclitols may be derived from aminosugars

52 via a yet to be determined mechanism (Sakuda, Sugiyama et al. 2001; Sugiyama, Nagasawa et al.

2002; Kudo, Kasama et al. 2007). Because of the biological roles of the aminocyclitols, the biosynthetic pathways of these natural products are under investigation with the hope that this knowledge could lead to the development of clinically significant therapeutics.

The Aminoglycosides

The aminoglycosides were the first important therapeutic agents produced by bacterial fermentation and all work through essentially the same mode of antibacterial action. They prevent the translation of proteins by binding the 30S subunit of the 70S prokaryotic ribosome and interfere specifically with transfer RNA interactions with messenger RNA in the decoding site of the ribosome (Davies 2006). Despite the similar antibacterial action, new studies using high resolution NMR analyses and X-ray crystallographic structures clearly describe differences in binding of the various classes of aminoglycosides and distinct modes of action (Lynch,

Gonzalez et al. 2003; Ogle and Ramakrishnan 2005). Examples amongst the aminocyclitol containing aminoglycosides are the antibiotics streptomycin, hygromycin, butirosin, neomycin, gentamycin (Figure 3.2).

Streptomycin was first isolated in 1943 by the group of S. A. Waksman and is historically the second great antibiotic after penicillin. Streptomycin was isolated from actinomycetes, the focus of Waksman’s career, and it was proven to be particularly effective in the treatment of tuberculosis, whereas penicillin is ineffective. It was his discovery of streptomycin that was the primary factor in the awarding of his Noble prize in medicine in 1952 (Kingston 2004). The precursor of streptomycin is myo-inositol, an ubiquitous compound playing a role in many cellular and physiological processes in living organisms. Myo-inositol is derivatized from glucose 6-phosphate in a process involving the enzyme 1-myo-inositol 1-phosphate synthase.

Figure 3.2. The aminoglycosides. Shown are the chemical structures of various antibiotics belonging to the aminoglycoside class of natural products

Myo-inositol is then tailored by a series of reactions to a more complex aminocyclitol (Mahmud

2009). In streptomycin biosynthesis, myo-inositol is oxidized to scyllo-inosose by myo-inositol dehydrogenase (StrI) and then transaminated to scyllo-inosamine by scyllo-inosose aminotransferase (StsC) to produce the aminocyclitol core of streptomycin (Mahmud, Flatt et al.

2007).

Hygromycin is another example of an aminoglycoside antibiotic which uses myo-inositol as a biosynthetic precursor (Mahmud 2009). Hygromycin was first isolated from Streptomyces hygroscopicus in 1953, and was first described to have a relatively broad spectrum of modest activity against gram-negative and gram-positive bacteria (Mann, Gale et al. 1953; Pittenger,

Wolfe et al. 1953). It has recently received a renewed interest due to its effectiveness in the treatment of swine dysentery by the anaerobic spirochete Serpulina hyodysenteriae (Omura,

Nakagawa et al. 1987). The tailoring of the aminocyclitol unit of this antibiotic involves the oxidation of C-5 of myo-inositol to give myo-inosose-5, catalyzed by a putative myo-inositol dehydrogenase, Hyg17, followed by the conversion to 2-L-2-amino-2-deoxyneo-inositol by the

PLP-dependent aminotransferase, Hyg8 (Habib, Scarsdale et al. 2003).

The precursory core of several aminoglycoside antibiotics such as butirosin, neomycin and gentamycin are formed by the 2-deoxy-scyllo-inosose (DOI) synthases, which catalyze the conversion of glucose 6-phosphate to DOI (Nango, Kumasaka et al. 2008; Mahmud 2009).

Furthermore, despite being synthesized by different genera of bacteria, these aminoglycosides also share the amino-pseudo-disaccharide paromamine as a common intermediate. The significant conservation of biosynthetic gene clusters required for paromamine biosynthesis facilitated the initial prediction of their biosynthetic pathways. In the synthesis of the paromamine intermediate, DOI is converted to 2-deoxy-scyllo-inosamine by a PLP-dependent

55 aminotransferase. Interestingly, this PLP-dependent aminotransferase is also responsible for the conversion of 2-deoxy-3-keto-scyllo-inosamine to 2-deoxystreptamine. A dehydrogenase oxidizes 2-deoxy-scyllo-inosamine to 2-deoxy-3-keto-scylloinosamine. N- acetylglucosaminyltransferase catalyzes the conversion of 2-deoxystreptamine to 2´-N- acetylparomamine. Finally, the paromamine core is completed by 2´-N-acetylparomamine deacetylase. Considering the number of homologous enzymes, the structural diversity of this particular set of aminoglycosides is remarkable. Glycosyltransferases further contribute to the diversity by mediating the attachment of various sugar moieties (Mahmud 2009).

The Five-Membered Ring Aminocyclitols

The five-membered ring aminocyclitols or aminocyclopentitols such as pactamycin, trehazolin, and allosamidin are of particular interest due to their ability to inhibit glycosidases and other carbohydrate recognizing enzymes by mimicking the corresponding substrates (Figure

3.3) (Winchester and Fleet 1992). Until recently, the only information available about the biosynthesis of aminocyclopentitols was derived from conventional feeding experiments with isotopically labeled precursors which identified glucose as the source of the aminocyclitol component (Weller and Rinehart 1978; Rinehart, Weller et al. 1980). Recently, the biosynthetic gene cluster of pactamycin was identified and analyses of the biosynthetic pathway of this aminocyclopentitol are underway (Kudo, Kasama et al. 2007; Ito, Roongsawang et al. 2009). It has been proposed that this unique aminocyclitol core is formed by a radical-mediated rearrangement of a glucosaminyl moiety by a Fe-S radical SAM oxidoreductase (Kudo, Kasama et al. 2007; Ito, Roongsawang et al. 2009).

The antibacterial pactamycin was isolated in 1962 from the soil bacterium Streptomyces pactum by scientists at the former Upjohn Company (Bhuyan 1962). This inhibitor of the 30S

Figure 3.3. The aminocyclopentitols. Shown are the chemical structures of various antibiotics belonging to the aminocyclopentitol class of natural products bacterial ribosome unit also showed strong anti-tumor activity but is too toxic for clinical use

(Bhuyan 1962; Hanessian, Vakiti et al. 2011). Pactamycin is comprised of a five-membered ring aminocyclitol unit linked to 6-methylsalicylic acid, 3-aminoacetophenone, and a 1,1- dimethylurea (Wiley, Jahnke et al. 1970; Rinehart, Weller et al. 1980). Hanessian, Vakiti et al. described pactamycin as the most densely functionalized naturally occurring aminocyclopentitol known (Hanessian, Vakiti et al. 2011).

In 1986, allosamidin was identified as an inhibitor of chitinase, an enzyme required for the proper degradation of chitin in insect cuticle. Such an agent capable of preventing the

57 degradation of old exoskeleton was sought as a potentially new biological insecticide. Isolated from the mycelial extract of Streptomyces sp. No. 1713, allosamidin is a pseudotrisaccharide constructed of two -linked N-acetyl-2-amino-2-deoxy-D-allopyranoside units with an aminocyclopentitol moiety attached at the reducing end (Sakuda, Isogai et al. 1986; Sakuda,

Sugiyama et al. 2001).

A 1991 screen for trehalase inhibitors uncovered a pseudodisaccharide termed trehazolin.

Trehazolin was isolated from a culture broth of Micromonospora sp. SANK 62390 and consists of an -glucose monomer linked to the aminocyclopentitol trehazolin via an isourea functionality. Trehazolin is a particularly potent trehalase inhibitor (Ando, Satake et al. 1991;

Masson, Philouze et al. 2005).

The C7N Aminocyclitols

The C7N aminocyclitols are a class of microbially-derived secondary metabolites, particularly from soil bacteria of the order of Actinomycetes (Mahmud 2003). Their biological activities are known to be brought about by an unsaturated aminocyclitol moiety, valienamine, contained within their chemical structures (Shick and Dunlap 2002). Although, some of the class members instead possess the epoxy, hydroxy or dihydro forms of valienamine (Mahmud 2003).

The common precursor to these metabolites is 2-epi-5-epi-valiolone, which is generated from sedoheptulose 7-phosphate by the sugar phosphate cyclase, 2-epi-5-epi-valiolone synthase

(Mahmud 2009). Examples of C7N aminocyclitols include acarbose, salbostatin, cetoniacytone, and validamycin (Figure 3.4) (Mahmud 2003). These members can be grouped based on the stage of biosynthesis in which intermediate phosphorylation comes into play. In the acarbose and salbostatin pathways, 2-epi-5-epi-valiolone is phosphorylated by a cyclitol kinase to 2-epi-5- epi-valiolone 7-phosphate (Zhang, Stratmann et al. 2002). Alternatively, in validamycin and

Figure 3.4. The C7N aminocyclitols. Shown are the chemical structures of various antibiotics belonging to the C7N aminocyclitols class of natural products

cetoniacytone biosynthesis, 2-epi-5-epi-valiolone is not phosphorylated, but is instead first epimerized to 5-epi-valiolone by a class of metalloproteins that appear to be a new type of α- hydroxyketo-epimerases belonging to the Vicinal Oxygen Chelate (VOC) superfamily (Wu, Flatt et al. 2007).

The C7N aminocyclitol family member, acarbose, is a pseudooligosaccharide isolated from Actinoplanes, which contains an unsaturated cyclitol unit and a 4-amino-4,6- dideoxyglucose unit. Acarbose potently inhibits sucrase, maltase, dextinase, and glycoamlyase activity while also showing some inhibition activity against amylose degrading enzymes

(Truscheit, Frommer et al. 1981). This clinically important α-glucosidase inhibitor has been proven useful in the treatment of type II insulin independent diabetes by preventing digestion of complex carbohydrates (Puls, Keup et al. 1977; Schmidt, Frommer et al. 1977). In September

1995 after years of availability in Europe, acarbose was approved by the FDA for use in the

United States as monotherapy in patients with non-insulin-dependent diabetes mellitus

(NIDDM).(Campbell, White et al. 1996)

Salbostatin is a basic, non-reducing pseudodisaccharide isolated from a fermentation culture polyether antibiotic salinomycin producer, Streptomyces albus ATCC 21838. It consists of valienamine linked to 2-amino-1,5-anhydro-2-deoxyglucitol (Vertesy, Fehlhaber et al. 1994).

Although salbostatin shows no antibiotic activity, it does effectively inhibit trehalase from porcine kidney (Mahmud 2003).

Cetoniacytone is a metabolite isolated from an endosymbiotic Actinomyces sp. strain Lu

9419 that lives in the intestines of the rose chaffer beetle (Cetonia aurata) (Schlorke, Krastel et al. 2002). The valienamine derived moiety of this unusual C7N aminocyclitol family member contains an alkylated amino group at the C-2 position instead of an acetylated amino group at the

C-1 position as in most family members. The moiety also contains an epoxide. Despite the structural anomalies, the core unit of cetoniacytone is derived via a product of the pentose phosphate pathway, sedoheptulose 7-phosphate (Wu, Flatt et al. 2009).

Validamycin was first identified during the course of screening for new antibiotics at the

Takeda Chemical Company. Isolated from the fermentation culture of Streptomyces hygroscopicus subsp. limoneus, this weakly basic, water-soluble antibiotic exhibited remarkable therapeutic effects against plant diseases (Iwasa, Yamamoto et al. 1970; Iwasa, Higashide et al.

1971). The biosynthesis of validamycin is of key interest in this document and is discussed in greater detail in later sections.

Validamycin A

Early Uses of Validamycin A

Validamycin A is a pseudotrisaccharide found within certain species of Streptomyces hygroscopicus and consists of valienamine and validamine connected via a nitrogen bridge. The second cyclitol also carries a glucose monomer attached via a glycosidic link (Mahmud, Lee et al. 2001). Validamycin A has long been used in the Far East as crop-protectant against rice- sheath blight and the damping-off of cucumber seedlings caused by the pathogenic fungi

Rhizoctonia solani and Pellicularia sasakii, respectively (Iwasa, Yamamoto et al. 1970; Iwasa,

Higashide et al. 1971). The antifungal activity of validamycin A is brought about by its ability to inhibit the enzyme trehalase. Trehalose is a disaccharide used chiefly within fungi as a carbohydrate storage compound comprising about 8% of the dry weight of the biomass. The administration of validamycin A results is an accumulation of toxic levels of trehalose

(Shigemoto, Okuno et al. 1992). This is observed as an abnormal branching of the tips of the main hyphae and severe repression of the growing fungi (Shibata, Uyeda et al. 1981).

Validamycin has also been reported to have an adverse reaction on insects which is presumably due to the similar prevalence of trehalose within some insects (Kameda, Asano et al. 1987). In addition, the validamycin derivative, voglibose, is widely used as a treatment for diabetes (Horii,

Fukase et al. 1986). As a natural product with known biological importance, validamycin and its derivatives are of great interest in agricultural and clinical research (Mahmud 2003).

Discovery of the Validamycin A Gene Cluster

Acarbose is a compound used as a treatment for type II insulin-independent diabetes.

Acarbose works by preventing the proper catabolism of complex carbohydrates through the inhibition of -glucosidases (Puls, Keup et al. 1977). A structural comparison to validamycin A revealed that validamycin A and acarbose share an identical C7N aminocyclitol moiety, valienamine (Mahmud 2003). The valienamine moiety of acarbose is known to originate from the cyclization of sedoheptulose 7-phosphate to 2-epi-5-epi-valionone by AcbC, an enzyme closely related to 3-dehydro-quinate synthases. Yu, Bai, et al. used the gene encoding AcbC

(abcC) within acarbose biosynthesis as a probe for genes encoding similar cyclization enzymes within the genome of S. hygroscopicus subsp. limoneus (Yu, Bai et al. 2005). The localization of the homologous gene valA allowed for the sequencing and analysis of a 45kb DNA fragment containing the entire validamycin A biosynthetic gene cluster (Bai, Li et al. 2006). Homologous validamycin A biosynthetic gene clusters have now been identified in two subspecies of

Streptomyces hygroscopicus, subsp. limoneus (vld) and subsp. jinggangensis (val) (Bai, Li et al.

2006; Singh, Seo et al. 2006). With the discovery of the validamycin A gene cluster, a mechanistic analysis of validamycin A biosynthesis could begin in earnest.

Biosynthetic Pathway of Validamycin A

Despite the biosynthetic pathway not being elucidated in its entirety, the formation of validamycin A by S. hygroscopicus is quite complicated. Both cyclitol components of validamycin A are derivatives of 2-epi-5-epi-valionone, the common precursor to the various

C7N-aminocyclitol-containing natural products (Figure 3.5). 2-epi-5-epi-valionone is generated by the cyclization of sedoheptulose 7-phosphate by the sugar phosphate cyclase, 2-epi-5-epi- valiolone synthase ValA (VldA), a homolog of AcbC (Bai, Li et al. 2006). Within validamycin

Figure 3.5. Early validamycin biosynthesis. Shown are the early enzymatic steps in the biosynthesis of validamycin. (DH? Represents a currently unknown dehydrogenase)

A biosynthesis 2-epi-5-epi-valionone is then epimerized to 5-epi-valionone by ValD, a new type of -hydroxyketo-epimerase belonging to the Vicinal Oxygen Chelate (VOC) superfamily (Bai,

Li et al. 2006; Xu, Zhang et al. 2009). Next is the dehydration of 5-epi-valionone to valienone by a yet to be determined dehydratase. At this point, the editing of the two cyclitols diverges.

ValC (VldC), a C7-cyclitol kinase, phosphorylates valienone to form valienone 7- phosphate (Figure 3.6) (Minagawa, Zhang et al. 2007). The conversion of valienone 7- phosphate to valienol 1-phosphate involves a stereoselective reduction of C-1, possibly by ValK, followed by the transfer of phosphate from C-7 to C-1. The latter step may be catalyzed by a phosphohexomutase, possibly ValO (Yang, Xu et al. 2011). Valienol 1-phosphate is

Nucleotidylated by ValB (VldB), a homolog of the sugar 1-phosphate nucleotidyltransferases

(Yang, Xu et al. 2011). The result is GDP-valienol, the first reactant in the in the condensation reaction forming the precursory validoxylamine A 7´-phosphate.

In the production of validamine 7-phosphate, the second reactant of the condensation reaction, valienone is most likely saturated by ValN (Figure 3.6). Although inconclusive, studies show that inactivation of the putative cyclitol reductase gene valN prevents the synthesis of validamycin A but instead produces unsaturated derivatives (Xu, Yang et al. 2009). The resulting validone is then phosphorylated. The C7-cyclitol kinase, VldC (ValC), is also responsible for the phosphorylation of this intermediate (Minagawa, Zhang et al. 2007).

Downstream, validone 7-phosphate may be recognized by the aminotransferase, ValM, to produce validamine 7-phosphate (Bai, Li et al. 2006).

The pseudo-glycosyltransferase VldE(ValL) is responsible for the condensation of GDP- valienol and validamine 7-phosphate via a non-glycosidic nitrogen bridge to form validoxylamine A 7´-phosphate (Figure 3.6) (Asamizu, Yang et al. 2011). The dephosphorylation of validoxylamine A 7´-phosphate by VldH is necessary before the addition of a glucose monomer by ValG (VldK) via a glycosidic linkage. ValG uses UDP-glucose as the source for the carbohydrate (Bai, Li et al. 2006; Xu, Minagawa et al. 2008). The final addition of the glucose monomer yields the natural antibiotic validamycin A.

Figure 3.6. Late validamycin biosynthesis. Shown are the late enzymatic steps in the biosynthesis of validamycin.

Glycosyltransferases

A Large and Diverse Class of Enzymes

Within the biosynthetic pathway of validamycin, the enzyme VldE is of particular interest because it mediates the condensation of the two aminocyclitol moieties via an amide linkage. A sequential and functional analysis relates VldE to a family of enzymes known as the glycosyltransferases (Bai, Li et al. 2006; Singh, Seo et al. 2006). The glycosyltransferases are an extremely large and diverse class of enzymes found in every kingdom of life and in all cellular compartments. Typically, they transfer a sugar residue from an activated donor to an acceptor.

The donor group is often a nucleotide sugar but there are examples of lipid phosphates and sugar-1-phosphates (Hansen, Bettler et al. 2010). The nucleotide sugar –dependent glycosyltransferases are often referred to as Leloir enzymes in honor of the 1970 Nobel Prize laureate Luis F. Leloir for his contributions to glycobiology (Lairson, Henrissat et al. 2008). The group of acceptors is also quite diverse, and the products of such reactions are oligo/polysaccharides, glycoproteins, glycolipids, and many other glycosylated natural products

(Hansen, Bettler et al. 2010). The obvious abundance of this class of enzymes is seen in the classification of glycosyltransferases by the CAZy database. The CAZy database divides the

91,000+ genes into 94 enzyme families (as of June 2012) (www.cazy.org) (Hansen, Bettler et al.

2010). In addition to classification based on amino acid sequence similarities, the glycosyltransferases are generally distinguished by their three-dimensional folds which consist primarily of Rossman-like // domains in either a GT-A or GT-B configuration. The GT-A or

GT-B folds are distinct from a sometimes associated transmembrane domain (Kikuchi and

Narimatsu 2006). Glycosyltransferases are further classified by the two possible stereochemical outcomes for the formed glycosidic bond. The anomeric configuration of the donated moiety is

66 either maintained or inverted in the product (Lairson, Henrissat et al. 2008). Recently, glycosyltransferases not adopting a GT-A or GT-B topology have been reported, and sequence searches have proposed a potential third topology, GT-C (Liu and Mushegian 2003; Lovering, de

Castro et al. 2007; Igura, Maita et al. 2008; Lizak, Gerber et al. 2011). As these cases are fairly uncommon and as a complete verification of GT-C glycosyltransferases by experimental means has not been performed (Lairson, Henrissat et al. 2008), the discussion of these “non-Leloir” glycosyltransferases within this document will end here.

GT-A and GT-B Topologies

Unlike the diverse representatives of the 110 families of the glycosidases, only two folds have been associated with the solved structures of the activated sugar-dependent glycosyltransferases, GT-A and GT-B, with uncharacterized families also predicted to adopt one of these two folds (Lairson, Henrissat et al. 2008). The GT-A fold, which consists of two abutting // domains forming a single continuous core -sheet, was first described in SpsA from Bacillus subtilis (Figure 3.7) (Charnock and Davies 1999).

SpsA is a glycosyltransferase implicated in the spore coat polysaccharide biosynthesis of

Bacillus subtilis (Stragier and Losick 1996). The N-terminal domain of SpsA consists of four parallel -strands and two -helices forming a nucleotide-binding fold. The C-terminal domain consists of a mixed -sheet flanked by three -helices forming an open groove which is the likely binding site for the acceptor group (Charnock and Davies 1999). Because of the abutting nature of the two Rossman-like domains, glycosyltransferases of the GT-A topology are often considered as a single domain fold (Figure 3.7A). Additionally, most GT-A glycosyltransferases contain an Asp-X-Asp signature responsible the coordination of the phosphate atoms of

Figure 3.7. Glycosyltransferase topologies. Shown are ribbon diagrams representing the crystallographic structures of enzymes demonstrating the two topologies associated with all glycosyltransferases. (A) SpsA is of the GT-A topology while (B) -glucosyltransferase (GTB) from T4-bacteriophage is of the GT-B topology.

68 nucleotide donors through the coordination of a divalent cation, Mg2+ or Mn2+ (Hansen, Bettler et al. 2010).

In comparison, the GT-B topology associated with glycosyltransferases features two

Rossman-like// domains oriented in an opposing, non-abutting manner. GT-B enzymes also lack a coordinated divalent cation. There exists a strong conservation of the C-terminal

Rossman-like domain of GT-B enzymes, which has been associated with the binding of nucleotides (Hansen, Bettler et al. 2010). The prototypical representative of the GT-B glycosyltransferases is -glucosyltransferase (GTB) from T4-bacteriophage (Hansen, Bettler et al. 2010). GTB is responsible for the glycosylation of doubled stranded DNA through transfer of glucose from UDP-glucose to the hydroxymethyl substituents of modified cytosine bases as a defense mechanism to prevent genome degradation by phage and host nucleases (Kornberg,

Kornberg et al. 1961). Structural studies revealed GTB to consist of two domains with similar topologies with the active site located within the cleft separating the two domains. The N- terminal domains consisted of a seven stranded parallel twisted -sheet and seven -helices while the C-terminal, nucleotide binding domain consists of a six stranded parallel twisted - sheet and five -helices. The catalytic site is located at the interface of the two domains (Figure

3.7B) (Vrielink, Ruger et al. 1994).

Inverting Glycosyltransferases

Glycosyltransferases are further classified by the retention or inversion of the anomeric configuration of the product with respect to the donor substrate. However there is no correlation between the anomeric state of the reaction product and the fold topology as there are examples both retaining and inverting reactions are catalyzed by both GT-A and GT-B folds (Lairson,

Henrissat et al. 2008).

Figure 3.8. Catalytic mechanisms of glycosyltransferases. Shown are the various catalytic mechanisms associated with glycosyltransferases: (A) an inverting SN2 reaction, (B) a retaining double displacement SN2 reaction, and (C) a retaining SNi-like reaction.

The inversion of anomeric centers by glycosyltransferases works through a direct replacement SN2-like reaction where the nucleophilic acceptor group displaces the activated phosphate leaving group (Figure 3.8A). Such a mechanism can be directly inferred from the catalytic mechanism of glycosidases. As the mechanistic strategy of inverting glycosyltransferases is well understood, examinations of the catalytic mechanisms focus on the

70 identification of the base catalyst and the method used to facilitate departure of the phosphate leaving group (Lairson, Henrissat et al. 2008).

Inverting enzymes with a GT-A topology include SpsA, -1,4-galactosyltransferase, N- acetylglucosaminyltransferase I, -1,3-N-acetylglucosaminyltransferases, and glucuronyltransferases (Lairson, Henrissat et al. 2008). Structural studies of these enzymes show a conserved aspartate (or glutamate) within hydrogen bonding distance of the attacking hydroxyl of the acceptor group which acts as a general base (Charnock and Davies 1999; Pedersen,

Tsuchida et al. 2000; Unligil, Zhou et al. 2000; Kakuda, Shiba et al. 2004; Jinek, Chen et al.

2006; Ramakrishnan, Ramasamy et al. 2006; Yang, Boze et al. 2009). Biochemical analysis show that the divalent cation bound by most GT-A enzymes is necessary to facilitate the leaving group by electrostatically stabilizing the forming negative charges on the phosphates of the leaving group (Lairson, Henrissat et al. 2008). Although there are exceptions such as metal ion- independent sialyltransferases and β-1,6-GlcNAc transferases which use hydroxyls of basic amino acids to stabilize the leaving group (Pak, Arnoux et al. 2006; Chiu, Lairson et al. 2007)

GT-B enzymes are more diverse in their selected modes of catalyzing glycosyl group transfer (Lairson, Henrissat et al. 2008). The prototypical β-glucosyltransferase (BGT) from T4 bacteriophage is an inverting glycosyltransferase which uses aspartate as a general base with the leaving group facilitated by positively charged side-chains. BGT does make use of a divalent cation, but unlike GT-A enzymes, the cation is involved in product release not facilitation of the leaving group (Lariviere and Morera 2002). The glycosyl transferases involved in the biosynthesis of the peptide antibiotic vancomycin also use an aspartate as the catalytic base but the leaving group is facilitated using a helix dipole and interactions with side chain hydroxyl and imidazole groups (Mulichak, Losey et al. 2001; Mulichak, Losey et al. 2003; Mulichak, Lu et al.

2004). The terpene/flavonoid glycosyltransferase UGT71G1, the flavonoid glucosyltransferase

VvGTI, the macrolide glycosyltransferases OleD and OleI, the human drug-metabolizing glucuronyltransferases UGT2B7, and the bifunctional N- and O-glucosyltransferase from

Arabidopsis thaliana also use a similar metal ion-independent mechanism to facilitate leaving group departure but the catalytic base was determined to be a histidine that interacts with an adjacent carboxylate group (Shao, He et al. 2005; Offen, Martinez-Fleites et al. 2006; Bolam,

Roberts et al. 2007; Brazier-Hicks, Offen et al. 2007; Miley, Zielinska et al. 2007).

Retaining Glycosyltransferases

Similar to inverting glycosyltransferases, retaining glycosyltransferases are found within both the GT-A and GT-B topologies. Unlike inverting glycosyltransferases, the catalytic mechanism of retaining glycosyltransferases cannot be directly inferred from the reaction of glycosidases. Glycosidases use a double-displacement SN2 reaction which works through a covalently bound glycosyl-enzyme intermediate. In such a mechanism, the donor group undergoes a nucleophilic attack by a nucleophilic residue which would displace the leaving phosphate group and result in a glycosyl-enzyme intermediate with an inverted anomic center at the donated group. This would be followed by a nucleophilic attack by the acceptor group which would displace the nucleophilic residue and again invert the anomeric configuration of the donated group resulting in a net retention of anomeric configuration of the donated group within the product (Figure 3.8B) (Lairson, Henrissat et al. 2008).

The reaction mechanism of retaining glycosyltransferases is much less understood.

A double displacement mechanism has been implicated in some retaining glycosyltransferases.

For instance, the bovine galactosyltransferase, α3GalT, contains a carboxylate side-chain which is correctly disposed to act as the nucleophile through the glycosyl-enzyme intermediate can be

72 formed. Mutation of this residue abolishes activity but activity can be rescued by azide

(Monegal and Planas 2006; Jamaluddin, Tumbale et al. 2007). However, most retaining glycosyltransferases lack a candidate for this role in close proximity to the anomeric reaction center and there is no strictly conserved signature within glycosyltransferase catalytic site to suggest such a mechanism. Therefore two possibilities exist; either the catalytic site undergoes a reorganization to provide an appropriate nucleophile candidate for a double displacement reaction, or retaining glycosyltransferases work through an alternative mechanism (Lairson,

Henrissat et al. 2008).

A proposed alternative is a SNi-like single nucleophilic substitution (Figure 3.8C). A SNi reaction is a special case SN1 reaction in which there is a partial decomposition of the donor group generating partial yet discrete ion pairs. Within glycosyltransferase reactions, this ion pair would be a partial negative charge on the leaving phosphate oxygen and a partial positive charge on the anomeric carbon. This short-lived transition state would be completed by the association with the attacking nucleophile of the acceptor group on the front-face of the anomeric carbon.

The formation of this immediate species occurs at a rate faster than that of solvent attack or the reorganization required for the nucleophilic attack from the back-face of the anomeric carbon.

In such a mechanism, the leaving phosphate oxygen would also act as a general base to activate the attacking nucleophile as it is the only moiety appropriately positioned to do so (Lairson,

Henrissat et al. 2008). Substrate/product phosphates are known to act as general bases and have been implicated in the mechanisms of farnesyl diphosphate (FPP) synthases from Escherichia coli and Trypanosoma cruzi (Hosfield, Zhang et al. 2004; Gabelli, McLellan et al. 2006).

Although SNi reactions have been proposed for both GT-A and GT-B topologies, the facilitation of the leaving groups would occur through differing manners as seen in inverting

73 glycosyltransferases. Retaining GT-A enzymes such as galactosyltransferase LgtC from

Neisseria meningitidis or rabbit muscle glycogenin use a divalent cation coordinated with the

Asp-X-Asp signature to facilitate the leaving group by stabilization of the forming negative charges (Persson, Ly et al. 2001; Gibbons, Roach et al. 2002), while retaining GT-B enzymes such as rabbit muscle glycogen phosphorylase (rmGP) and E. coli α-(1,3) glucosyltransferase

WaaG use positive side chains to facilitate the leaving group (Green, Cori et al. 1942; Martinez-

Fleites, Proctor et al. 2006).

Oxocarbenium Ion-Like Transition State

Functional and sequential analysis of VldE suggests a mechanism analogous to that of trehalose 6-phosphate synthase (OtsA). OtsA has been associated with a SNi-like mechanism.

The retaining glycosyltransferase reaction is stabilized by an oxocarbenium ion-like transition state (Figure 3.8C) (Lairson, Henrissat et al. 2008). The aminocyclitol components of validamycin lack the bridging oxygen found within a sugar moiety. So, the oxocarbenium ion- like transition state is not possible within aminocyclitol condensation by VldE. Perhaps, the transition state of during VldE catalysis is stabilized by the olefinic moiety through a mechanism similar to the formation of an oxocarbenium ion within OtsA (Asamizu, Yang et al. 2011).

Hopefully, structural studies of VldE can reveal the validity of the proposed mechanism.

CHAPTER 4: MECHANISTIC INSIGHTS INTO VALIDOXYLAMINE A 7'- PHOSPHATE SYNTHESIS BY VLDE USING THE STRUCTURE OF THE ENTIRE PRODUCT COMPLEX

Introduction

Glycosyltransferases comprise one of the most numerous and diverse groups of enzymes in nature. They are responsible for the formation of oligo/polysaccharides, glycoproteins, glycolipids, and many other glycosylated natural products by transferring a sugar moiety from an activated donor sugar to a sugar (or non-sugar) acceptor. This abundant group of proteins consists of 92 families encoded by more than 83,400 genes.(Hansen, Bettler et al. 2010)

However, only a fraction of those genes has actually been functionally characterized. Our comparative bioinformatics studies suggest that among those reported as glycosyltransferases are also pseudo-glycosyltransferases (such as VldE), which do not recognize sugars as substrates but rather catalyze the formation of non-glycosidic C-N bonds in the biosynthesis of C7N- aminocyclitol-containing natural products such as acarbose and validamycin A (Figure 4.1).(Bai,

Li et al. 2006; Singh, Seo et al. 2006; Asamizu, Yang et al. 2011) Acarbose, an -glucosidase inhibitor, has been proven useful in the treatment of type II insulin-independent diabetes, whereas validamycin A, a natural trehalase inhibitor, is an antifungal antibiotic that has long been used to protect crops from soil borne diseases such as rice sheath blight and the dumping- off of cucumber seedlings.(Iwasa, Yamamoto et al. 1970; Iwasa, Higashide et al. 1971; Xia and

Jiao 1986; Mahmud 2003)

Validamycin A is a pseudo-trisaccharide whose structure is comprised of validoxylamine

A and glucose (Figure 4.1). The final step in validamycin A biosynthesis is the attachment of glucose to the precursory validoxylamine A by the action of the glycosyltransferase VldK

(ValG).(Bai, Li et al. 2006; Xu, Minagawa et al. 2008) Validoxylamine A is generated through

Figure 4.1. The VldE and OtsA reactions. This figure was prepared with ChemBioDraw (ChemBioDraw Ultra 10.0 CambridgeSoft, 100 Cambridge Park Drive, Cambridge, MA 02140). The product and substrates of the VldE and OtsA catalyzed reactions are shown. Note the considerable similarity between the ligands of VldE and OtsA, and the conservation of the anomeric centers. The biosynthetic products of both the completed enzymatic pathways are also shown. VldE catalyzes the formation of validoxylamine A 7´-phosphate via a non-glycosidic C- N bond between GDP-valienol and validamine 7-phosphate. After the validoxylamine A 7´- phosphate has been produced; VldH and VldK complete the catalytic synthesis of Validamycin A. OtsA catalyzes the formation of trehalose 6-phosphate via a glycosidic bond between UDP- glucose and glucose 6-phosphate. OtsB dephosphorylates trehalose 6-phosphate to produce trehalose. the dephosphorylation of validoxylamine A 7´-phosphate by VldH while validoxylamine A 7´- phosphate (VDO) results from a condensation of GDP-valienol and validamine 7-phosphate

(both are pseudo-sugars) by the pseudo-glycosyltransferase, VldE.(Mahmud, Lee et al. 2001;

Wehmeier and Piepersberg 2004; Seo, Im et al. 2006; Singh, Seo et al. 2006; Flatt and Mahmud

2007; Mahmud 2009; Asamizu, Yang et al. 2011)

The mechanism by which non-glycosidic C-N bond is formed by a pseudo- glycosyltransferase is not entirely understood. However, because of the structural similarity of validoxylamine A 7´-phosphate to trehalose 6-phosphate (Figure 4.1), it has been speculated that the mechanism of the pseudo-glycosyltransferase VldE is similar to that of the glycosyltransferase, trehalose 6-phosphate synthase (OtsA).(Asamizu, Yang et al. 2011) VldE and Escherichia coli OtsA only share a modest 19% sequence identity (29% homology) (Figure

4.2), but they are both catalogued as members of the GT20 glycosyltransferase family by the

CAZy database (www.cazy.org).(Coutinho, Deleury et al. 2003) OtsA mediates the transfer of glucose moiety from UDP-glucose to glucose 6-phosphate to form trehalose 6-phosphate (Figure

4.1). Similar to VldE, the product of OtsA conserves the anomeric configuration of the donor moiety. Glycosyltransferases have been shown to both retain and invert the anomeric state of the carbon C-1 of the donor moiety. The inversion of the anomeric center by glycosyltransferases has been well explored and is known to be carried out by a simple nucleophilic substitution.

However, the net retention of the anomeric center is much more ambiguous, but it is thought to occur through either a double displacement (SN2 X2) or internal return mechanism

(SNi).(Lairson, Henrissat et al. 2008) For a double displacement nucleophilic substitution reaction to occur, a nucleophilic catalytic base must be available in close vicinity to the sugar anomeric carbon to form a covalent intermediate. OtsA apparently lacks such a residue in the catalytic site and analyses of theoretical energy manifolds along with recent studies of kinetic isotope effects have substantiated this unusual enzymatic reaction mechanism within

OtsA.(Gibson, Turkenburg et al. 2002; Tvaroska 2006; Lee, Hong et al. 2011) Lastly, structural

Figure 4.2. An alignment of VldE and OtsA. Shown is the protein sequence alignment of VldE and OtsA as generated by ClustalW2.(Larkin, Blackshields et al. 2007) Residues whose side-chains are involved in ligand binding are indicated with shaded boxes. Conserved residues are indicated with unfilled boxes. The two-dimensional secondary structure of VldE and OtsA are also illustrated next to the corresponding sequence with -helices represented by spirals and -strands represented by arrows. Numerical values for helices and strands are assigned.

78 studies of OtsA, using a bi-substrate inhibitor as a product mimic, place the atoms involved in catalysis in an orientation favorable for an internal return mechanism and show that the leaving phosphate acts as a general base to deprotonate the incoming nucleophile of the acceptor group.(Errey, Lee et al. 2010) The SNi mechanism of OtsA works through an oxocarbenium ion- like transition state.(Errey, Lee et al. 2010; Lee, Hong et al. 2011) However, no oxocarbenium ion formation is possible in the VldE-catalyzed non-glycosidic C-N coupling reaction.

Alternatively, it is proposed that the olefinic moiety of GDP-valienol may play a critical role in facilitating the coupling reaction.(Asamizu, Yang et al. 2011) Coupling reactions involving an allylic moiety have been demonstrated in other biosynthetic enzymes, e.g., farnesyl diphosphate

(FPP) synthases.(Kellogg and Poulter 1997; Thulasiram and Poulter 2006) However, in FPP synthases, a nucleophilic substitution reaction takes place at a carbon center with a diphosphate acting as a leaving group instead of nucleotidyl diphosphate. In addition, mechanistically FPP synthases adopt a stereospecific SN1 reaction with an inversion of the configuration, leaving the actual mechanism behind the unique catalytic function of the retaining VldE enzyme unclear.

Hopefully, structural studies would clarify whether or not a SNi mechanism is conserved within the retaining pseudo-glycosyltransferase, VldE.

Here, we report the three-dimensional structures of VldE in various liganded states using crystallographic techniques. The structure of VldE was solved by molecular replacement using the structure of OtsA as a search model. We have elucidated the structures of the unliganded

VldE, in complex with guanosine 5´-diphosphate (GDP), in complex with GDP and Trehalose

(TRE), and in complex with GDP and VDO. Similar to OtsA, VldE is comprised of two

Rossman // domains which are oriented in a GT-B configuration (Figure 4.3). The active site, which is located at the interface of the Rossman domains, and the selective interactions

Figure 4.3. The overall fold of VldE. All protein renderings were constructed using PyMOL (The PyMOL Molecular Graphics System, Version 1.2r3pre, Schrödinger, and LLC.). Shown is the overall fold of VldE as well as comparison to OtsA by superimposition. (A) The overall fold of the monomeric VldE is represented in a ribbon diagram. The monomer is rendered with the N-terminal domain in red, and the C-terminal domain in blue. The -helical C-terminus, which stretches back across both domains, is rendered in green. VldE consists of twin Rossman-like // domains in a GT-B configuration with the catalytic site marked by VDO and GDP at the interface of the two domains. (B) To compare the overall fold of VldE (green) to OtsA (gray), the folding patterns, which were represented by a tracing of C-alphas, were superimposed within a wall-eye stereo image. allowing for the binding of GDP but not uridine 5´-diphosphate (UDP) are described in our study

(Figure 4.4A-B). The crystallographic investigation of VldE while binding the complete product complex, VDO and GDP, supports the proposed conservation of a SNi catalytic mechanism analogous to the mechanism of OtsA. Both the “open” and “closed” conformations of the catalytic site are also described within this study.

Materials and Methods

Purification and Crystallization

VldE from S. hygroscopicus subsp. limoneus was expressed and purified with methods similar to those previously described.(Asamizu, Yang et al. 2011) 6XHis-tagged VldE was expressed in E. coli BL21 (DE3) pLysS and was purified using Ni-NTA affinity chromatography. Affinity

Figure 4.4. The catalytic site. Shown is a comparison of the VldE and OtsA catalytic sites in ribbon diagrams. Residues/molecules of interest are represented in stick models. The dotted lines mark significant interactions. The preferential binding of GDP by VldE is demonstrated by comparing (A) the protein-ligand interactions within the VldE active site (green) with (B) the protein-ligand interactions between OtsA and UDP within the OtsA active site (gray). The protein interactions with ribose and phosphate are conserved between VldE and OtsA. However, there are differing interactions with the nucleotide base groups. The large purine makes interactions with the residues Arg321, Asn323, Asn361, and Thr366. The ribose and phosphate moieties interact with Arg290, Lys295, Leu387, Ser388, and Glu391. Within OtsA, Leu344, Ile295, Pro297, and His338 only allow for the binding of the smaller pyrimidine. The ribose and phosphate moieties make interactions with residues Arg262, Lys267, Leu365, and Glu369. The mesh represents the |Fo| -|Fc| electron density omit map of the GDP binding site. The map is contoured at 3.0 levels. (C) Shown is a superimposition of the VldE cyclitol binding sites in the presence (yellow) and absence (green) of validoxylamine A 7´-phosphate. The mesh represents the |Fo| -|Fc| electron density omit map of the VDO binding site. The map is contoured at 3.0 levels. VDO makes interactions with residues the side-chains of residues Asp158, His182, Arg12, Asn325, Arg326, and Asp383. VDO also makes interactions with the backbones of residues 384-387. Binding of the acceptor cyclitol is recognized by conformation changes by the side-chains of residues Asp158, Tyr159, and Arg326. (D) Shown is a comparison by superimposition of VDO binding within the catalytic sites of VldE (yellow) and OtsA (gray). Note the strong conservation of residue and ligand positions.

82 chromatography was performed using 40mM Hepes buffer, pH 7.5 with 300mM NaCl, 20mM imidazole, 5mM -mercaptoethanol, and 10% glycerol. Elution from the Ni-NTA resin was achieved using the same buffer but with 175mM imidazole. Using dialysis, the affinity chromatography buffer was exchanged with VldE storage buffer consisting of 10mM Tris-HCl, pH 7.5 with 5% glycerol, 0.1mM dithiothreitol, and 1mM MgCl2. VldE was concentrated to

10mg/mL, and crystals were grown using sitting drop vapor diffusion.

In the case of VldE and VldE•GDP crystals, the mother liquor consisted of 100mM Tris-

HCl, pH 8.0 with 30-35% polyethylene glycol 3,350. GDP, at a concentration of 3mM, was added to protein stocks which were intended to grow VldE•GDP crystals by co-crystallization.

Crystals with a size of 0.1 X 0.1 X 0.5 mm were grown over a period of four to twelve weeks.

Crystallization took place at a temperature of 293K. Cryoprotectant solutions, consisting of artificial mother liquor enriched with 20% ethylene glycol, were added to crystal drops at a

1:1(v/v) ratio. In the case of VldE•GDP crystals, 5mM GDP was added to the cryoprotectant solution. Crystals were flash-cooled in liquid nitrogen after soaking in the cryoprotectant solution for 20-40 minutes.

In the case of VldE•GDP•VDO crystals, the mother liquor consisted of 100mM Tris-HCl, pH 8.0 with 25-30% polyethylene glycol 3,350. The drop was formed through a mixture of mother liquor, protein, and seed stock in a ratio of 1.6:2.0:0.4 (v/v). Seed stocks were prepared from crystal grown in similar conditions using Seed Beads (Hampton Research) in a stabilization buffer of 100mM Tris-HCl, pH 8.0 with 31% polyethylene glycol 3,350. The Crystals with a size of 0.1 X 0.1 X 0.5 mm were grown over a period of four to twelve weeks at a temperature of

293K. Cryoprotectant solutions, consisting of artificial mother liquor enriched with 20% ethylene glycol, were added to crystal drops at a 1:1(v/v) ratio. 5mM GDP and 10mM VDO

83 were added to the cryoprotectant solutions. Crystals were flash-cooled in liquid nitrogen after soaking in the cryoprotectant solution for 20-40 minutes.

In the case of VldE•GDP•TRE crystals, the mother liquor consisted of 50mM Tris-HCl, pH 8.0, 18-28% polyethylene glycol 3,350, 200-500mM NaCl, and 20mM MgCl2. VldE was kept at a concentration of 10mg/mL, and crystals were grown using sitting drop vapor diffusion with a 1:1(v/v) mixture of protein and mother liquor. Before the drops were set, 3% trehalose and 1mM GDP were added to the protein stocks. The Crystals with a size of 0.1 X 0.1 X 0.1 mm were grown over a period of four to eight weeks at a temperature of 293K. Cryoprotectant solutions, consisting of reservoir solution enriched with 20% ethylene glycol, 3% trehalose, and

1mM GDP were added to crystal drops at a 1:1(v/v) ratio. Crystals were flash-cooled in liquid nitrogen after soaking in the cryoprotectant solution for 20-40 minutes.

Data Collection and Processing

All diffraction data except for the VldE•GDP dataset was collected at the Northeastern

Collaborative Access Team (NE-CAT) beamline at the Advanced Photon Source (Argonne

National Laboratory). Data was recorded at 100K on an ADSC Q315 (315mm X 315mm) detector and processed by NE-CAT's RAPD automated processing

(https://rapd.nec.aps.anl.gov/rapd), which uses XDS (Kabsch 2010) for integration and scaling.

A 2.0Å VldE dataset was collected at a wavelength of 0.9479 Å while oscillating the crystal 0.5 degrees for each frame. A 2.11Å VldE•GDP•VDO dataset was collected at a wavelength of

0.9792 Å while oscillating the crystal 1.0 degrees for each frame. A 2.81Å VldE•GDP•TRE dataset was collected at a wavelength 0.9792Å while oscillating the crystal 1.0 degrees for each frame. The 2.15Å VldE•GDP dataset was collected at NSLS Beamline X6A at a wavelength

1.0Å while oscillating the crystal 1.0 degrees for each frame. X-ray data was collected on a

ADSC Quanta CCD detector at 100K and was integrated, merged, and scaled using

HKL2000.(Otwinowski and Minor 1997) Although the cell dimensions were similar, the unliganded VldE crystal belonged to the P2 space group, while the VldE•GDP and

VldE•GDP•VDO crystals belonged to the P21 space group. The unit cell dimensions of the

VldE•GDP•TRE dataset were dissimilar and belonged to the C2 space group. Diffraction data statistics are summarized in Table 4.1.

Structure Determination and Refinement

To determine the structure of VldE, molecular replacement (MR) was performed.

Generation of a search model from the OtsA structure (PDB accession code 1UQU (Gibson,

Tarling et al. 2004)) and molecular replacement was carried out using MrBUMP (Keegan and

Winn 2007) to orchestrate molecular replacement via the MOLREP (Vagin and Teplyakov 1997) module in the CCP4 (Collaborative Computational Project 1994) program suite. The resulting

MR model was used to build a partially sequenced model (~400 residues) using the

AUTOBUILD (Terwilliger 2004; Afonine, Grosse-Kunstleve et al. 2005; Terwilliger, Grosse-

Kunstleve et al. 2008; Terwilliger, Grosse-Kunstleve et al. 2008) function of the PHENIX

(Adams, Afonine et al. 2010) software suite. Further automated model building was carried out using the ARP/wARP (Langer, Cohen et al. 2008) module within the CCP4 (Collaborative

Computational Project 1994) software suite. The model was finished by manual building within

COOT (Emsley, Lohkamp et al. 2010). The model was refined using restrained refinement by

REFMAC5 (Vagin, Steiner et al. 2004) with TLS parameters defined by the TLSMD server.(Painter and Merritt 2006; Painter and Merritt 2006) Two monomers were modeled within the asymmetric unit, and non-crystallographic symmetry (NCS) restraints were used during refinement. Waters were incorporated into the models by referring to the |Fo|-|Fc| omit

Table 4.1. Statistics of reflection data and structure refinements (VldE). Liganding VldE VldE•GDP VldE•GDP•VDO VldE•GDP•TRE Space Group P2 P21 P21 C2 Unit Cell Dimension (Å) 84.75 X 48.43 X 47.82 X 121.15 47.96 X 120.71 320.82 X 122.82 122.66 X 84.33 X 84.24 X 93.86 Unit Cell -angles(°) 91.88 91.74 91.63 91.62 Wilson Plot B Value (Å2) 48.74 35.96 40.37 53.40 Resolution Range (Å) 48.92-1.95 37.52-2.15 47.94-2.11 49.15-2.81 Reflections Observed 245,918 368,911 205,530 220,052 Unique Reflections 69,317 51,924 55,049 88,243 Reflections Rfree Set 3,559 2,645 2,790 4,421 Completeness (%) 98.1 (91.8) 99.9 (100.0) 99.7 (99.0) 99.3 (98.5) Redundancy 3.5 (3.3) 7.1 (7.1) 3.7 (3.7) 2.5 (2.5) 13.0 (2.7) 25.2 (3.1) 16.5 (2.7) 12.2 (2.7) Rsym 0.045 (0.421) 0.078(0.522) 0.049(0.433) 0.058(0.304) Rwork 0.190 0.197 0.191 0.211 Rfree 0.222 0.209 0.222 0.258 No. TLS Bodies 6 n/a n/a n/a No. of Amino Acids 957 948 950 2,738 No. of Protein Atoms 7,474 7,378 7,361 21,796 No. of Hetero Atoms 0 58 120 214 No. of Waters 492 405 488 202 R.M.S.D Bond Lengths (Å) 0.022 0.012 0.017 0.016 Angles (°) 1.67 1.30 1.76 1.47 Mean B Factor 41.85 40.21 39.62 62.41 Protein Atoms (Å2) 42.63 40.19 39.59 62.61 Hetero Atoms (Å2) n/a 34.34 37.63 62.57 Water Atoms (Å2) 29.38 40.93 40.50 40.82 Ramachandran Outliers (%) 0.5 0.4 0.4 0.4 Ramachandran Favored (%) 97.8 97.8 97.3 95.5 Poor Rotamers (%) 1.6 1.3 1.6 1.5

Table 4.1 continued

Rsym = Σ h(Σ j|Ihj - |/ Σ Ih,j), where h=set of Miller indices, j=set of observations of reflection h, and =the mean intensity. R.M.S.D. values are deviation from ideal values. Rcrys= Σh||Fo,h| – |Fc,h||/ Σh|Fo,h|. Rfree was calculated using 5% of the complete data set excluded from refinement. The numbers in parentheses represent values from the highest resolution shell (2.08-2.00Å for VldE, 2.23-2.15Å for VldE•GDP, 2.22-2.11Å for VldE•GDP•VDO, and 2.96-2.81Å for VldE•GDP•TRE).

87 maps. Out of the possible 994 residues, 957 residues were modeled with 97.8% within the most favored regions of the Ramachandran plot. Although 0.5% are reported as outliers, they are all within well-defined electron densities. The final model, which lacked residues 1, 2, and 482-

497, has a Rwork of 0.190 and an Rfree of 0.222. The structure refinement statistics are summarized in Table 4.1.

The VldE•GDP, VldE•GDP•VDO, and VldE•GDP•TRE structures were solved using the unliganded VldE structure as a search model within the AUTOMR (Mccoy, Grosse-Kunstleve et al. 2007) function of the PHENIX (Adams, Afonine et al. 2010) software suite. The models were finished by manual building within COOT.(Emsley, Lohkamp et al. 2010) The models was refined the PHENIX.REFINE (Afonine, Grosse-Kunstleve et al. 2005) function of the PHENIX

(Adams, Afonine et al. 2010) software suite. Ligands and waters were incorporated into the models by referring to the |Fo|-|Fc| omit maps.

Within the VldE•GDP structure, two monomers were modeled within the asymmetric unit, and NCS restraints were used during refinement. Out of the possible 994 residues, 948 residues were modeled with 97.8% are within the most favored regions of the Ramachandran plot. Although 0.4% are reported as outliers, they are all within well-defined electron densities.

Within the VldE•GDP•TRE structure, six monomers were modeled within the asymmetric unit, and NCS restraints were implemented during refinement for polypeptide chain pairs A and D, B and E, and C and F. Out of the possible 2,982 residues, 2,738 residues were modeled with 95.5% within the most favored regions of the Ramachandran plot. Although 0.4% are reported as outliers, they are all within well-defined electron densities.

Within the VldE•GDP•VDO structure, two monomers were modeled within the asymmetric unit, and NCS restraints were used during refinement. Out of the possible 994

88 residues, 950 residues were modeled with 97.3% within the most favored regions of the

Ramachandran plot. Although 0.4% are reported as outliers, they are all within well-defined electron densities.

The Ramachandran outliers at residues Asp158 and Asn386 were common within all the monomers of all the models of VldE.

Diffraction and structure refinement statistics are summarized in Table 4.1. Coordinates and structure factors have been deposited in the Protein Data Bank with accession numbers:

VldE (PDB ID: 3VDM), VldE•GDP (PDB ID: 4F96), VldE•GDP•VDO (PDB ID: 4F97), and

VldE•GDP•TRE (PDB ID: 4F9F).

Results

Overall Structure

As was recently described in a crystallographic study of ValL (identical but for a four residue insertion) (Zheng, Zhou et al. 2012), a VldE homolog from S. hygroscopicus subsp. jinganggensis, VldE consists of twin Rossman // domains in the GT-B configuration

(Vrielink, Ruger et al. 1994) (Figure 4.3A) with the terminal helix (16) extending back across the C-terminal Rossman-like domain to reach the N-terminal Rossman-like domain with a turn at

Trp458 as it crosses the interface of the two domains. Despite a modest 19% sequence identity

(29% homology) (Figure 4.2), the overall fold of VldE is remarkably similar to OtsA (Figure

4.3B), a fellow member of the GT20 family. Unlike the previous study however, the core of the

N-terminal domain of our apo enzyme is found with the catalytic site in an “open” conformation instead of “closed”. In this “open” conformation, the core -sheet of the N-terminal Rossman- like domain consists of only ten strands instead of twelve. Two strands described as part of the core sheet in the previous study now are observed in a unique -hairpin motif (2 and 3)

89 extending away from the catalytic site. Simultaneously, there is a dramatic shift of residues 33-

47 away from the catalytic center. Unfortunately, coordinates from the structural study of ValL were not publicly available at the time of this study, so a complete comparison could not be performed. However, the closed conformation of VldE will be described in later sections of this article using our own experimental data.

The GDP bound structure of VldE (VldE•GDP) described here is also in an “open” conformation and the binding of GDP did not have an effect on the global conformation of the apo enzyme (R.M.S.D. of C = 0.350). Surprisingly, the structure of VldE in complex with the entirety of its products (VldE•GDP•VDO), GDP and VDO was also in an open conformation and a comparison to the VldE•GDP model showed that the binding of VDO also did not have an effect of the global conformation of VldE (R.M.S.D. C = 0.262).

Lastly, we also elucidated the structure of VldE in complex with both GDP and TRE

(VldE•GDP•TRE). Within the VldE•GDP•TRE structure, six monomers were modeled within the asymmetric unit (ASU). The six monomers of the ASU assumed three alternative conformers. The monomers of the first conformation contained both GDP and trehalose, and this first conformer was found to also have an “open” catalytic site with similar global conformation to the VldE•GDP•VDO complex (R.M.S.D. C = 0.705). The last two conformers within the

VldE•GDP•TRE structure had GDP but not trehalose bound, and these two conformers contained a “closed” catalytic site as described in the previous studies.(Zheng, Zhou et al. 2012) These alternative conformations, which have significant differences when compared the open

VldE•GDP structure also described here (R.M.S.D. C = 2.304, R.M.S.D. C = 2.395), are further discussed in later sections.

The Nucleotide Binding Site

Due to the conservation of key residues within their binding pockets, the binding of ligands within VldE are thought to be analogous to OtsA.(Gibson, Turkenburg et al. 2002) Here we provided experimental support for the analogous binding of GDP within VldE and UDP within

OtsA. (Figure 4.4A) Within VldE, the ribose and phosphate moieties of GDP make interactions with the side-chains of the residues Arg290, Lys 295 and Glu391; as well as with the backbone atoms of residues Leu387 and Ser388 (Figure 4.5A). These interactions are conserved within

OtsA (Figure 4.4B). The -phosphate of GDP does make a single unique interaction with

Ser388 within VldE. This interaction is not possible within OtsA as there is a valine at the same position. Nonetheless, the ribose and phosphate moieties of GDP within the VldE catalytic pocket are near perfectly superimposable onto the ribose and phosphate moieties of UDP within the OtsA catalytic pocket. The conserved position of the -phosphate of GDP is critically important because the bond between this phosphate and the donated cyclitol is broken during the transfer to the acceptor. As will be described later in this study, the -phosphate of GDP maintains an interaction with the anomeric carbon of validoxylamine A 7´-phosphate in the complete product complex (VldE•GDP•VDO). This interaction is similarly found within OtsA and similarly provides strong evidence in support of an SNi mechanism.(Lee, Hong et al. 2011)

The guanine moiety within the VldE•GDP complex makes interactions with the hydrophilic residues Arg321, Asn323, and Thr366 (Figure 4.5A). A comparison to the binding of a uracil moiety of the nucleotide activated donor sugar within OtsA allows an immediate understanding of the preferential binding of the guanine moiety by VldE. The preference of guanine by VldE is mediated by these three hydrophilic residues, while the preference for a uracil moiety is mediated by three hydrophobic residues at the same positions. Within OtsA, the

Figure 4.5. Ligand interactions. Shown is a ChemBioDraw figure showing the hydrogen bond network between VldE and (A) GDP and then (B) VDO.

92 uracil is surrounded by the hydrophobic residues Ile295, Pro297, and Leu344. This small hydrophobic binding pocket facilitates the binding of a uracil moiety. Interactions between VldE and GDP are summarized in Table 4.2.

Cyclitol Binding Sites

Provided within this article is the first structure of VldE in complex with the true products of its catalyzed reaction, VDO and GDP (Figure 4.4C). The interactions between GDP and VldE are described within the previous section, but here the interactions between VDO and

VldE are shown to be analogous to the interactions previously described by the OtsA•VDO complex.(Errey, Lee et al. 2010) The hydroxyls of the donated cyclitol make numerous interactions with the backbone amides of VldE at residues Gly384, Gln385, Asn386, and

Leu387. The donated cyclitol also makes interactions with the side-chains of residues His182 and Asp383 (Figure 4.5B). The interactions by donated cyclitol are conserved within the catalytic site of OtsA (Figure 4.4D). Also conserved is the interaction of the -phosphate with the anomeric carbon of the donated cyclitol. As in OtsA, the phosphate of the leaving nucleotide is involved in the stabilization of the transition state during the concerted, front-faced nucleophilic attack of a SNi reaction, and at a distance of 2.6Å from the attacking nucleophile, the phosphate can act as a general base to deprotonate the amine.(Errey, Lee et al. 2010)

The binding of the acceptor cyclitol within VldE is recognized by a shift in side-chain conformations within the catalytic site. Asp158 swings about 62-83° towards the catalytic center in recognition of the hydroxyls of the acceptor cyclitol. Upon the binding of the phosphorylated acceptor group, the guanidine moiety of Arg326 enters into an interaction with the phosphate oxygens. This interaction allows Tyr159 to swing about 110-112° to make an interaction with

Arg326. The acceptor cyclitol also interacts with the side-chain of Asn325. These interactions

Table 4.2. Notable interactions between VldE and ligands. Validoxylamine A GDP Trehalose 7´-Phosphate Ribose 5’- Donor Acceptor Donor Acceptor Protein Atom Guanine phosphate Cyclitol Cyclitol-P Sugar Sugar Arg321* NH2 O6 (3.3) Asn323* ND2 N1 (3.3) ND2 N2 (3.4) Asp360 O O6 (3.3) Asn361 OD1 N1 (2.6) OD1 N2 (2.9) Asp362 N O6 (2.9) O O6 (3.2) Thr366* OG1 N7 (2.6) OG1 O6 (3.6) Arg290 NE O2B (2.8) NH2 O1B (2.9) Lys295 NZ O3A (3.1) NZ O1B (3.0) Leu387 N O1A (3.4) N O2A (2.8) Ser388 OG O1A (2.6) N O1A (3.1) Glu391 OE1 O2’ (2.5) OE2 O3’ (2.6) His182 ND1 OAQ (2.8) O6P (3.7) Asp383 OD1 OAS (2.9) O3P (3.3) Gly384 N OAS (3.0) O3P (2.6) Gln385 N OAS (2.7) O3P (2.2) N OAR (2.8) O4P (3.2) Asn386 N OAS (3.4) O3P (3.4) N OAR (2.7) O4P (2.6) Leu387 N OAR (3.6) O4P (3.7)

Table 4.2 continued. Asp158 OD1 OAO (3.3) OD2 OAP (2.3) Asn325 ND2 OAX (2.4) OD1 OAX (3.6) Arg326 NH1 OAW (3.0) NE OAY (3.6) Arg290 NE O4 (3.7) Values in parentheses are distances given in angstroms. Donor/Acceptor is an indicator of relative position within the catalytic site. Distances describing GDP interactions with VldE are taken from the VldE•GDP model. * Indicates residues not conserved in OtsA that make guanine specific interactions.

95 are conserved within OtsA except at Asn325. At the same position within OtsA there is Ser299 which does not make an interaction with VDO (Figure 4.4D).(Asamizu, Yang et al. 2011)

Despite the conservation of the interactions between the catalytic site and VDO, there is no observed conformational closing of the catalytic site upon the binding of the acceptor cyclitol as was witnessed in structural studies of OtsA.(Gibson, Turkenburg et al. 2002; Gibson, Tarling et al. 2004; Errey, Lee et al. 2010) However, a closed conformation was described in the recently published structural studies of ValL.(Zheng, Zhou et al. 2012) In later sections, such a conformational shift will be described within our structural studies of VldE, but this shift was not triggered by the binding of the acceptor cyclitol.

We were also able to elicit the structure of VldE in complex with both GDP and trehalose

(VldE•GDP•TRE). (Figure 4.6A) Within this model, GDP maintains the interactions previously described elsewhere within this study. The binding pose of trehalose is similar to what was described by the recent structural study of VldE.(Zheng, Zhou et al. 2012) The sugar moiety of trehalose within donor cyclitol binding site makes interactions with the backbone amides of residues Gly384, Gln385, Asp386, and Leu387 as well with the side-chains of Asp383 and

His182. Unsurprisingly, this conserves the interactions between the enzyme and the structurally similar donor cyclitol as was described in the VldE•GDP•VDO model. However, the sugar moiety of trehalose occupying the acceptor cyclitol binding site makes only a single significant interaction with Arg290 in our study. A superimposition of trehalose and VDO within the VldE catalytic site shows that trehalose does not assume a binding pose comparable to VDO (Figure

4.6B). This is most likely due to the absence of a phosphorylated ligand. Within the

VldE•GDP•VDO model, Arg326 and Asn325 interact with the phosphorylated cyclitol and most likely help to ensure the proper docking of validamine 7-phosphate for catalysis. Indicatively,

Figure 4.6. Trehalose within the catalytic site. Shown is a trehalose (TRE) within the VldE cyclitol binding site in ribbon diagrams. Residues/molecules of interest are represented in stick models. The dotted lines mark significant interactions. (A) The mesh represents the |Fo| -|Fc| electron density omit map of trehalose within the VldE catalytic site (pink). The map is contoured at 3.0 levels. Trehalose makes interactions with the backbone amides of residues Gly384, Gln385, Asp386, and Leu387 as well with the side-chains of Asp383, His182 and Arg290. (B) Shown is a superimposition of the catalytic sites of VldE•GDP•VDO model (yellow) and the VldE•GDP•TRE model (pink) in ribbon diagrams. Trehalose does not assume a binding pose comparable to VDO. This is most likely to do the absence of a phosphorylated ligand as Arg326 and Asn325 swing out of the catalytic site.

Arg325 and Asn326 swing out of the catalytic site within our trehalose bound structure.

Interactions between VldE and VDO and TRE are summarized in Table 4.2.

Observed Changes in Conformation

While solving the VldE•GDP•TRE structure, six monomers of VldE were placed within the asymmetric unit. Represented within the six monomers were three differing conformations of the N-terminal Rossman-like domain (two monomers per conformation). (Figure 4.7A-C)

The first conformer had GDP and trehalose bound and is described in the previous section. This conformer is quite similar to the other models with “open” catalytic sites expect for residues 325-328 which swing out of the catalytic site upon the binding of trehalose. The 97

Figure 4.7. Conformational changes. Shown are ribbon diagrams of the three conformers of VldE modeled using the VldE•GDP•TRE crystallographic data. Residues/molecules of interest are represented in stick models. Arrows indicate the direction of residue movement. Ligands bound within each conformer are shown to provide a point of reference for comparison. (A-C) The global conformation of each conformer (gray) is shown with the areas of conformational change highlighted by coloring (pink, blue, cyan). (D-F) Residues 11-50 for each conformer is shown. Note that this region of residues is capable of 10.9Å shift towards the catalytic center and that strands of the -hairpin motif (2 and 3) move simultaneously with strands 6 and 5 to extend the core -sheet from ten to twelve strands. (G-I) A view of helix 3 for each conformer is shown. Note that residues at the kink of the helix are capable of reorganizing.

99 second and third conformers have only GDP within the catalytic site and both of these conformers share a dramatic shift in residues 11-50 which result in the “closing” of the catalytic site (Figure 4.7D-F). Resultantly, Gly33, Gly34, and Thr35 move 10.9Å towards the catalytic center, and within 4Å of the ligand atoms. Additionally, helix , which originally consisted of residues 36-41, coils further to include residues 42-47. This shift occurs concurrently with the rotation of the -hairpin motif (2 and 3) unique to VldE. Additionally, the last two of ten strands (6 and 5) comprising the core -sheet of the N-terminal Rossman-like // domain move towards the compacting catalytic site. The movements of the unique -hairpin motif and the two strands of the core -sheet narrow the distance between them and allow the necessary interactions to unite them into a single twelve-stranded core -sheet. The “closed” conformation described here is similar to what was described in the structural study of

ValL.(Zheng, Zhou et al. 2012)

A conformational change which resulted in the closing of the catalytic site was described within the structural studies of OtsA. Residues were observed to make a >10Å movement towards the catalytic center upon binding of the accepting sugar in OtsA.(Gibson, Turkenburg et al. 2002; Gibson, Tarling et al. 2004) The conformational shift was thought to be due to the recognition of the phosphate moiety of the accepting glucose 6-phosphate by Arg9. However, with the lack of an acceptor group within the catalytic site, the conserved arginine at position 12 of VldE cannot be responsible for the closing of the catalytic site. Therefore, the closing of the

VldE catalytic site is more likely to work at least partially through a more dynamic, equilibrium based mechanism.

An additional conformation change was also observed within the third conformer found within the VldE•GDP•TRE crystal (Figure 4.7G-I). Within the 3, there is a kink at residues 96-

100

102. Within the third conformer, the kinked residues shift to form a more organized secondary structure. Again, the observed conformational change is not concerted with the binding of ligand and further supports the notion that VldE moves through equilibrium based conformations.

Conformational change at this position was observed in the previous structural studies, and similarly the change seems to be focused around Gln96.(Zheng, Zhou et al. 2012)

Discussion

Mechanistic Considerations

The net retention of the anomeric center is thought to occur through either a double displacement (SN2 X2) or internal return mechanism (SNi).(Lairson, Henrissat et al. 2008)

Recent evidence substantiates claims that the conservation of the anomeric center configuration within OtsA is due to an internal return mechanism.(Errey, Lee et al. 2010; Lee, Hong et al.

2011) If the reaction of VldE were to alternatively go through a double displacement nucleophilic substitution mechanism, a residue must be available to make a covalent intermediate. Comparatively, the hydrolysis of glycosides is known to occur through a nucleophilic substitution involving a covalent intermediate.(Tews, Perrakis et al. 1996; Vocadlo,

Davies et al. 2001) The most likely candidate residue within VldE would be a nucleophilic

His182. As can be seen in Figure 4.4D, this residue is also conserved within the catalytic site of

OtsA (His154). At a 5.1Å from the anomeric center, His182 is not observed to be immediately available for nucleophilic substitution in our structural studies, and as of yet there has been no significant, direct evidence to support a mechanism by which the non-glycosidic coupling reaction of VldE occurs through a covalent intermediate.

Our structural evidence demonstrates a strict conservation of nucleotide and cyclitol orientation within VldE and OtsA and suggests that the glycosyltransferase reaction of OtsA and

101 the pseudo-glycosyltransferase reaction of VldE do occur through similar, concerted but asynchronous internal return mechanisms. Therefore, the olefinic moiety of GDP-valienol must play a critical role in facilitating the coupling reaction through a mechanism similar to the formation of an oxocarbenium ion within OtsA as was previously suggested (Asamizu, Yang et al. 2011). Through such a mechanism, the transition state is formed by the coordination of the allylic carbon on donated cyclitol, the bridging amine of the acceptor cyclitol-phosphate and the leaving oxygen of the donor diphosphate-nucleotide. As in OtsA (Errey, Lee et al. 2010), the leaving phosphate is in a position to deprotonate the incoming nucleophile of the acceptor cyclitol (Figure 4.8A-B).

Supportively, a distance matrix search using DALI (Holm and Rosenstrom 2010) revealed a strong similarity between VldE and OtsA, cetyl--D-Glucosaminyl L-Malate

Synthase (BaBshA), 3-Phospho-1-D-I-Inosityl-2-Acetamido-2-Deoxy--D-Glucopyranoside

Synthase (MshA), Trehalose Synthase (TreT), Phosphatidylinositol Mannosyltransferase

(PimA), and Glycogen Synthase (GlgA). (Table 4.3) All are retaining glycosyltransferases that have been associated with internal return mechanisms.(Horcajada, Guinovart et al. 2006;

Vetting, Frantom et al. 2008; Batt, Jabeen et al. 2010; Errey, Lee et al. 2010; Parsonage, Newton et al. 2010; Woo, Ryu et al. 2010) (Table 4.3)

Significance

The biosynthetic pathways of natural products exhibiting antibiotic-like properties have recently become of great interest. As seen in the discovery of the antibacterial erythromycin, the anthelmintic avermectins, and the antitumor indolocarbozes, the genetic manipulation of biosynthetic gene clusters along with the use of alternative biosynthetic precursors, and recombinant proteins for chemoenzymatic synthesis can result in the generation of an entire

102

Figure 4.8. The SNi mechanism of VldE. Shown are ChemBioDraw figures comparing the SNi mechanism of (A) OtsA to the proposed SNi mechanism of (B) VldE. The olefinic moiety of GDP-valienol plays a critical role in facilitating the coupling reaction through a mechanism similar to the formation of an oxocarbenium ion within OtsA. Both transition states are formed by the coordination of the allylic carbon on the donated group, the bridging nucleophile of the acceptor group and the leaving oxygen of the donor diphosphate-nucleotide.

Table 4.3. DALI server results Enzyme Name R.M.S.D Trehalose-Phosphate Synthase (OtsA) 2.4 cetyl--D-Glucosaminyl L-Malate Synthase 2.9 (BaBshA)

3-Phospho-1-D-I-Inosityl-2-Acetamido-2-Deoxy-- 3.7 D-Glucopyranoside Synthase (MshA) Trehalose Synthase (TreT) 3.1 Phosphatidylinositol Mannosyltransferase (PimA) 3.1 Glycogen Synthase (GlgA) 3.1 - Listed in order of DALI ranking

103 array of biologically active compounds.(McDaniel, Thamchaipenet et al. 1999; Cropp, Wilson et al. 2000; Sanchez, Mendez et al. 2006; Mahmud, Flatt et al. 2007) The gene cluster responsible for the biosynthesis of validamycin A has only been recently discovered, this study and others are underway to elucidate the individual processes within this biosynthetic pathway of this C7N- aminocyclitol.(Bai, Li et al. 2006)

The C7N-aminocyclitols (e.g., acarbose, validamycin, cetoniacytone, and salbostatin) belong to a class of natural products which include the aminoglycosides (e.g., streptomycin, hygromycin, butirosin, and neomycin), and the five membered aminocyclitols or cyclopentitols

(e.g., pactamycin, trehazolin, and allosamidin). The aminoglycosides are a diverse group of natural products which were amongst the first clinical antibiotics. The C7N-aminocyclitols are an even more diverse group of natural products with an entire range of biological activities.(Mahmud 2009) A better understanding of the biosynthesis of C7N-aminocyclitols at the molecular level could allow for the generation of novel, natural product analogs which could include an array of structurally altered antibiotics. The line of inquiry provided within this article hopes to add to that understanding.

104

CHAPTER 5: CONCLUSIONS

As it is a technique that allows for the elucidation of structural information of a target molecule at the atomic level, protein crystallography is an extremely powerful tool. The three- dimensional graphics of proteins produced by crystallographers allows for the illustration of bonding networks and the differentiation of closely related isoenzymes. Crystallographic studies can also explain substrate specificity and provide insight, support, or validation to the underlying chemistry of a catalytic mechanism. This work describes the use of x-ray crystallography in exactly such a manner. Protein crystallography was put to use in the examination of the protein enzymes, PFKB3 and VldE.

The bifunctional enzyme is solely responsible for the cellular concentrations of an effector of glucose metabolism, fructose-2,6-bisphosphate. The inducible isoform has been implicated in the “Warburg Effect” in cancer biology and is unique because of a high bisphosphatase/kinase ratio. However, the low bisphosphatase activity of PFKFB3 allowed for a unique opportunity to study the bisphosphatase reaction. Although the experiments described within this work are not the first crystallographic studies of PFKB3 (Kim, Manes et al. 2006;

Kim, Cavalier et al. 2007), much insight was still obtained from this study. As previously described the degradation of fructose-2,6-bisphosphate is mediated through a mechanism typical of members of the histidine phosphatase family (Rigden 2008). The reaction begins with the nucleophilic attack by a catalytic histidine which forms a phospho-histidine intermediate. The residue side-chains forming the phosphate pocket stay in contact with the phosphate oxygens during the entirety of the reaction. There is a linear transfer of phosphate to the histidine and a catalytic side-chain acts as a general acid/base in the facilitation of the leaving fructose-6- phosphate and in the activation of a water molecule for the degradation of the phospho-histidine

105 intermediate. This study provided the last experimental data necessary to provide a complete visualization of catalysis from binding of substrate within the Michaelis complex through the transition states to the release of products. Also reported is a novel conformational change in the

C-terminal residues which suggests a role in substrate/product binding /release. Hopefully these observations could provide insight to other researchers investigating other histidine phosphatase family members.

The crystallographic studies of VldE reveal an enzyme with a global fold typical of GT-B glycosyltransferases that markedly generates a non-glycosidic bond. Furthermore, catalysis seems to be mediated by an internal return mechanism that has been associated with many glycosyltransferases which allows for the retention of the anomeric configuration of the donated moiety (Lairson, Henrissat et al. 2008). The preference of this glycosyltransferase family member for GDP as the activating nucleotide is clearly explained by the selective hydrogen bond network within the catalytic site. These structural studies provide the next step in the understanding of the validamycin biosynthetic pathway and can provide insight into the synthesis of other C7N containing natural products. Additionally, understanding of the molecular basis of catalysis by VldE could directly lead to efforts of scaffold diversification of validamycin for pharmacophoric optimization of anti-biotics with potential clinical significance through the genetic manipulation of the biosynthetic gene cluster, the use of alternative biosynthetic precursors, or the use of recombinant proteins.

106

REFERENCES Adams, P. D., P. V. Afonine, et al. (2010). "PHENIX: a comprehensive Python-based system for macromolecular structure solution." Acta Crystallographica Section D-Biological Crystallography 66: 213-221.

Afonine, P. V., R. W. Grosse-Kunstleve, et al. (2005). "The Phenix refinement framework." CCP4 Newsletter(July): Contribution 8.

Akinaga, S., K. Sugiyama, et al. (2000). "UCN-01 (7-hydroxystaurosporine) and other indolocarbazole compounds: a new generation of anti-cancer agents for the new century?" Anticancer Drug Design 15(1): 43-52.

Algaier, J. and K. Uyeda (1988). "Molecular cloning, sequence analysis, and expression of a human liver cDNA coding for fructose-6-P,2-kinase:fructose-2,6-bisphosphatase." Biochemical and Biophysical Research Communications 153(1): 328-333.

Ando, O., H. Satake, et al. (1991). "Trehazolin, a New Trehalase Inhibitor." Journal of Antibiotics 44(10): 1165-1168.

Asamizu, S., J. T. Yang, et al. (2011). "Pseudoglycosyltransferase Catalyzes Nonglycosidic C-N Coupling in Validamycin A Biosynthesis." Journal of the American Chemical Society 133(31): 12124-12135.

Atsumi, T., J. Chesney, et al. (2002). "High expression of inducible 6-phosphofructo-2- kinase/fructose-2,6-bisphosphatase (iPFK-2; PFKFB3) in human cancers." Cancer Research 62(20): 5881-5887.

Bai, L., L. Li, et al. (2006). "Functional analysis of the validamycin biosynthetic gene cluster and engineered production of validoxylamine A." Chemistry & Biology 13(4): 387-397.

Bailey, S. (1994). "The Ccp4 Suite - Programs for Protein Crystallography." Acta Crystallographica Section D-Biological Crystallography 50: 760-763.

Batt, S. M., T. Jabeen, et al. (2010). "Acceptor Substrate Discrimination in Phosphatidyl-myo- inositol Mannoside Synthesis: Structural and Mutational Analysis of Mannosyltransferase Corynebacterium Glutamicum PimB." Journal of Biological Chemistry 285(48): 37741- 37752.

Bauer, D. E., M. H. Harris, et al. (2004). "Cytokine stimulation of aerobic glycolysis in hematopoietic cells exceeds proliferative demand." Faseb Journal 18(9): 1303-1305.

Bazan, J. F., R. J. Fletterick, et al. (1989). "Evolution of a Bifunctional Enzyme - 6- Phosphofructo-2-Kinase Fructose-2,6-Bisphosphatase." Proceedings of the National Academy of Sciences of the United States of America 86(24): 9642-9646.

107

Bhuyan, B. K. (1962). "Pactamycin Production by Streptomyces Pactum." Applied Microbiology 10(4): 302-&.

Bing, F. C. (1971). "The history of the word 'metabolism'." Journal of the History of Medicine and Allied Sciences 26(2): 158-180.

Bodansky, O. (1972). "Acid phosphatase." Advances in Clinical Chemistry 15: 43-147.

Bolam, D. N., S. Roberts, et al. (2007). "The crystal structure of two macrolide glycosyltransferases provides a blueprint for host cell antibiotic immunity." Proceedings of the National Academy of Sciences of the United States of America 104(13): 5336- 5341.

Brahimi-Horn, C. and J. Pouyssegur (2006). "The role of the hypoxia-inducible factor in tumor metabolism growth and invasion." Bull Cancer 93(8): E73-80.

Brazier-Hicks, M., W. A. Offen, et al. (2007). "Characterization and engineering of the bifunctional N- and O-glucosyltransferase involved in xenobiotic metabolism in plants." Proceedings of the National Academy of Sciences of the United States of America 104(51): 20238-20243.

Bui, T. and C. B. Thompson (2006). "Cancer's sweet tooth." Cancer Cell 9(6): 419-420.

Calvo, M. N., R. Bartrons, et al. (2006). "PFKFB3 gene silencing decreases glycolysis, induces cell-cycle delay and inhibits anchorage-independent growth in HeLa cells." Febs Letters 580(13): 3308-3314.

Campbell, L. K., J. R. White, et al. (1996). "Acarbose: Its role in the treatment of diabetes mellitus." Annals of Pharmacotherapy 30(11): 1255-1262.

Campbell, W. C. and G. W. Benz (1984). "Ivermectin - a Review of Efficacy and Safety." Journal of Veterinary Pharmacology and Therapeutics 7(1): 1-16.

Caro, J. (2001). "Hypoxia regulation of gene transcription." High Altitude Medicine & Biology 2(2): 145-154.

Charnock, S. J. and G. J. Davies (1999). "Structure of the nucleotide-diphospho-sugar transferase, SpsA from Bacillus subtilis, in native and nucleotide-complexed forms." Biochemistry 38(20): 6380-6385.

Cheek, S., H. Zhang, et al. (2002). "Sequence and structure classification of kinases." Journal of Molecular Biology 320(4): 855-881.

Chesney, J., R. Mitchell, et al. (1999). "An inducible gene product for 6-phosphofructo-2-kinase with an AU-rich instability element: role in tumor cell glycolysis and the Warburg

108

effect." Proceedings of the National Academy of Sciences of the United States of America 96(6): 3047-3052.

Chesney, J., S. Telang, et al. (2005). "Targeted disruption of inducible 6-phosphofructo-2-kinase results in embryonic lethality." Biochemical and Biophysical Research Communications 331(1): 139-146.

Chi, H. B., X. N. Yang, et al. (2000). "Targeted deletion of Minpp1 provides new insight into the activity of multiple inositol polyphosphate phosphatase in vivo." Molecular and Cellular Biology 20(17): 6496-6507.

Chiu, C. P. C., L. L. Lairson, et al. (2007). "Structural analysis of the alpha-2,3-sialyltransferase cst-i from Campylobacter jejuni in apo and substrate-analogue bound forms." Biochemistry 46(24): 7196-7204.

Chu, D. T. W. (1995). "Section Review Anti-infectives: Recent developments in 14- and 15- membered macrolides." Expert Opinion on Investigational Drugs 4(2): 65-94.

Claus, T. H., M. R. El-Maghrabi, et al. (1984). "The role of fructose 2,6-bisphosphate in the regulation of carbohydrate metabolism." Current Topics in Cellular Regulation 23: 57-86.

Clem, B., S. Telang, et al. (2008). "Small-molecule inhibition of 6-phosphofructo-2-kinase activity suppresses glycolytic flux and tumor growth." Molecular Cancer Therapeutics 7(1): 110-120.

Coleman, D. E., A. M. Berghuis, et al. (1994). "Structures of active conformations of Gi alpha 1 and the mechanism of GTP hydrolysis." Science 265(5177): 1405-1412.

Collaborative Computational Project, N. (1994). "The Ccp4 Suite - Programs for Protein Crystallography." Acta Crystallographica Section D-Biological Crystallography 50: 760- 763.

Colomer, D., J. L. Vivescorrons, et al. (1987). "Control of Phosphofructokinase by Fructose 2,6- Bisphosphate in Lymphocytes-B and B-Chronic Lymphocytic-Leukemia Cells." Cancer Research 47(7): 1859-1862.

Cortes, J., S. F. Haydock, et al. (1990). "An Unusually Large Multifunctional Polypeptide in the Erythromycin-Producing Polyketide Synthase of Saccharopolyspora-Erythraea." Nature 348(6297): 176-178.

Coutinho, P. M., E. Deleury, et al. (2003). "An evolving hierarchical family classification for glycosyltransferases." Journal of Molecular Biology 328(2): 307-317.

Crepin, K. M., M. I. Darville, et al. (1989). "Characterization of Distinct Messenger-Rnas Coding for Putative Isozymes of 6-Phosphofructo-2-Kinase/Fructose-2,6- Bisphosphatase." European Journal of Biochemistry 183(2): 433-440.

109

Cropp, T. A., D. J. Wilson, et al. (2000). "Identification of a cyclohexylcarbonyl CoA biosynthetic gene cluster and application in the production of doramectin." Nature Biotechnology 18(9): 980-983.

Dang, C. V. and G. L. Semenza (1999). "Oncogenic alterations of metabolism." Trends in Biochemical Sciences 24(2): 68-72.

Danishefsky, S. (2010). "On the potential of natural products in the discovery of pharma leads: a case for reassessment." Natural Product Reports 27(8): 1114-1116.

Darville, M. I., M. Chikri, et al. (1991). "A Rat Gene Encoding Heart 6-Phosphofructo-2-Kinase Fructose-2,6-Bisphosphatase." Febs Letters 288(1-2): 91-94.

Darville, M. I., K. M. Crepin, et al. (1989). "5' Flanking Sequence and Structure of a Gene Encoding Rat 6-Phosphofructo-2-Kinase Fructose-2,6-Bisphosphatase." Proceedings of the National Academy of Sciences of the United States of America 86(17): 6543-6547.

Davies, J. E. (2006). "Aminoglycosides: ancient and modern." The Journal of Antibiotics (Tokyo) 59(9): 529-532.

DeBerardinis, R. J., J. J. Lum, et al. (2008). "The biology of cancer: Metabolic reprogramming fuels cell growth and proliferation." Cell Metabolism 7(1): 11-20.

Denoya, C. D., R. W. Fedechko, et al. (1995). "A second branched-chain alpha-keto acid dehydrogenase gene cluster (bkdFGH) from Streptomyces avermitilis: its relationship to avermectin biosynthesis and the construction of a bkdF mutant suitable for the production of novel antiparasitic avermectins." Journal of Bacteriology 177(12): 3504-3511.

Deprez, J., D. Vertommen, et al. (1997). "Phosphorylation and activation of heart 6- phosphofructo-2-kinase by protein kinase B and other protein kinases of the insulin signaling cascades." Journal of Biological Chemistry 272(28): 17269-17275.

Donadio, S., M. J. Staver, et al. (1991). "Modular Organization of Genes Required for Complex Polyketide Biosynthesis." Science 252(5006): 675-679.

Dubos, R. J. (1951). Louis Pasteur, free lance of science. London,, Gollancz.

Dutton, C. J., S. P. Gibson, et al. (1991). "Novel Avermectins Produced by Mutational Biosynthesis." Journal of Antibiotics 44(3): 357-365.

Eknoyan, G. (1999). "Santorio Sanctorius(1561-1636) - Founding father of metabolic balance studies." American Journal of Nephrology 19(2): 226-233.

El-maghrabi, M. R., T. H. Claus, et al. (1982). "Regulation of Rat-Liver Fructose 2,6- Bisphosphatase." Journal of Biological Chemistry 257(13): 7603-7607.

110

El-maghrabi, M. R., T. H. Claus, et al. (1982). "Regulation of 6-Phosphofructo-2-Kinase Activity by Cyclic Amp-Dependent Phosphorylation." Proceedings of the National Academy of Sciences of the United States of America-Biological Sciences 79(2): 315- 319.

El-Maghrabi, M. R., F. Noto, et al. (2001). "6-phosphofructo-2-kinase/fructose-2,6- bisphosphatase: suiting structure to need, in a family of tissue-specific enzymes." Current Opinion in Clinical Nutrition & Metabolic Care 4(5): 411-418.

Emsley, P. and K. Cowtan (2004). "Coot: model-building tools for molecular graphics." Acta Crystallographica Section D-Biological Crystallography 60: 2126-2132.

Emsley, P., B. Lohkamp, et al. (2010). "Features and development of Coot." Acta Crystallographica Section D-Biological Crystallography 66: 486-501.

Errey, J. C., S. S. Lee, et al. (2010). "Mechanistic Insight into Enzymatic Glycosyl Transfer with Retention of Configuration through Analysis of Glycomimetic Inhibitors." Angewandte Chemie-International Edition 49(7): 1234-1237.

Fischer, C., L. Rodriguez, et al. (2002). "Digitoxosyltetracenomycin C and glucosyltetracenomycin C, two novel elloramycin analogues obtained by exploring the sugar donor substrate specificity of glycosyltransferase ElmGT." Journal of Natural Products 65(11): 1685-1689.

Fisher, A. J., C. A. Smith, et al. (1995). "X-ray structures of the myosin motor domain of Dictyostelium discoideum complexed with MgADP.BeFx and MgADP.AlF4." Biochemistry 34(28): 8960-8972.

Flatt, P. M. and T. Mahmud (2007). "Biosynthesis of aminocyclitol-aminoglycoside antibiotics and related compounds." Natural Product Reports 24(2): 358-392.

Fothergillgilmore, L. A. and H. C. Watson (1989). "The Phosphoglycerate Mutases." Advances in Enzymology and Related Areas of Molecular Biology 62: 227-313.

Furuya, E. and K. Uyeda (1980). "An Activation Factor of Liver Phosphofructokinase." Proceedings of the National Academy of Sciences of the United States of America- Biological Sciences 77(10): 5861-5864.

Furuya, E. and K. Uyeda (1981). "A Novel Enzyme Catalyzes the Synthesis of Activation Factor from Atp and D-Fructose-6-P." Journal of Biological Chemistry 256(14): 7109-7112.

Furuya, E., K. Uyeda, et al. (1981). "The Effect of Synthetic and Natural Activation Factor on Phosphofructokinase." Federation Proceedings 40(6): 1605-1605.

111

Furuya, E., M. Yokoyama, et al. (1982). "An Enzyme That Catalyzes Hydrolysis of Fructose- 2,6-Bisphosphate." Biochemical and Biophysical Research Communications 105(1): 264- 270.

Furuya, E., M. Yokoyama, et al. (1982). "Regulation of Fructose-6-Phosphate 2-Kinase by Phosphorylation and Dephosphorylation - Possible Mechanism for Coordinated Control of Glycolysis and Glycogenolysis." Proceedings of the National Academy of Sciences of the United States of America-Biological Sciences 79(2): 325-329.

Gabelli, S. B., J. S. McLellan, et al. (2006). "Structure and mechanism of the farnesyl diphosphate synthase from Trypanosoma cruza: Implications for drug design." Proteins- Structure Function and Bioinformatics 62(1): 80-88.

Gatenby, R. A. and R. J. Gillies (2004). "Why do cancers have high aerobic glycolysis?" Nature Reviews Cancer 4(11): 891-899.

Gibbons, B. J., P. J. Roach, et al. (2002). "Crystal structure of the autocatalytic initiator of glycogen biosynthesis, glycogenin." Journal of Molecular Biology 319(2): 463-477.

Gibson, R. P., C. A. Tarling, et al. (2004). "The donor subsite of trehalose-6-phosphate synthase - Binary complexes with UDP-glucose and UDP-2-deoxy-2-fluoro-glucose at 2 angstrom resolution." Journal of Biological Chemistry 279(3): 1950-1955.

Gibson, R. P., J. P. Turkenburg, et al. (2002). "Insights into trehalose synthesis provided by the structure of the retaining glucosyltransferase OtsA." Chemistry & Biology 9(12): 1337- 1346.

Golicnik, M. (2010). "Metallic Fluoride Complexes as Phosphate Analogues for Structural and Mechanistic Studies of Phosphoryl Group Transfer Enzymes." Acta Chimica Slovenica 57(2): 272-287.

Goren, N., A. Manzano, et al. (2000). "6-Phosphofructo-2-kinase/fructose-2,6-bisphosphatase expression in rat brain during development." Molecular Brain Research 75(1): 138-142.

Green, A. A., G. T. Cori, et al. (1942). "Crystalline muscle phosphorylase." Journal of Biological Chemistry 142(1): 447-448.

Gribble, G. W., Berthel, S. J. (1993). A survey of indolo[2,3-a]carbazoles and related natural products. Studies in Natural Product Chemistry. A. U. Rahman. Amsterdam, Elsevier. 12: 365-409.

Habib, E. S. E., J. N. Scarsdale, et al. (2003). "Biosynthetic origin of hygromycin A." Antimicrob Agents Chemother 47(7): 2065-2071.

112

Hafner, E. W., B. W. Holley, et al. (1991). "Branched-Chain Fatty-Acid Requirement for Avermectin Production by a Mutant of Streptomyces-Avermitilis Lacking Branched- Chain 2-Oxo Acid Dehydrogenase-Activity." Journal of Antibiotics 44(3): 349-356.

Hamilton, J. A., M. J. Callaghan, et al. (1997). "Identification of PRG1, a novel progestin- responsive gene with sequence homology to 6-phosphofructo-2-kinase/fructose-2,6- bisphosphatase." Journal of Molecular Endocrinology 11(4): 490-502.

Hanessian, S., R. R. Vakiti, et al. (2011). "Total Synthesis of Pactamycin." Angewandte Chemie- International Edition 50(15): 3497-3500.

Hansen, S. F., E. Bettler, et al. (2010). "Exploring genomes for glycosyltransferases." Molecular Biosystems 6(10): 1773-1781.

Hasemann, C. A., E. S. Istvan, et al. (1996). "The crystal structure of the bifunctional enzyme 6- phosphofructo-2-kinase/fructose-2,6-bisphosphatase reveals distinct domain homologies." Structure 4(9): 1017-1029.

Heine-Suner, D., M. A. Diaz-Guillen, et al. (1998). "Sequence and structure of the human 6- phosphofructo-2-kinase/fructose-2,6-bisphosphatase heart isoform gene (PFKFB2)." European Journal of Biochemistry 254(1): 103-110.

Hirata, T., M. Watanabe, et al. (2000). "Inhibition of tumor cell growth by a specific 6- phosphofructo-2-kinase inhibitor, N-bromoacetylethanolamine phosphate, and its analogues." Bioscience Biotechnology and Biochemistry 64(10): 2047-2052.

Holm, L. and P. Rosenstrom (2010). "Dali server: conservation mapping in 3D." Nucleic Acids Research 38: W545-W549.

Horcajada, C., J. J. Guinovart, et al. (2006). "Crystal structure of an archaeal glycogen synthase - Insights into oligomerization and substrate binding of eukaryotic glycogen synthases." Journal of Biological Chemistry 281(5): 2923-2931.

Horii, S., H. Fukase, et al. (1986). "Synthesis and alpha-D-glucosidase inhibitory activity of N- substituted valiolamine derivatives as potential oral antidiabetic agents." Journal of Medicinal Chemistry 29(6): 1038-1046.

Hosfield, D. J., Y. M. Zhang, et al. (2004). "Structural basis for bisphosphonate-mediated inhibition of isoprenoid biosynthesis." Journal of Biological Chemistry 279(10): 8526- 8529.

Huang, F. L., S. F. Haydock, et al. (2005). "The neomycin biosynthetic gene cluster of Streptomyces fradiae NCIMB 8233: characterisation of an aminotransferase involved in the formation of 2-deoxystreptamine." Organic & Biomolecular Chemistry 3(8): 1410- 1418.

113

Huang, L. E., Z. Arany, et al. (1996). "Activation of hypoxia-inducible transcription factor depends primarily upon redox-sensitive stabilization of its alpha subunit." Journal of Biological Chemistry 271(50): 32253-32259.

Hue, L. and G. G. Rousseau (1993). "Fructose 2,6-bisphosphate and the control of glycolysis by growth factors, tumor promoters and oncogenes." Advances in Enzyme Regulation 33: 97-110.

Igura, M., N. Maita, et al. (2008). "Structure-guided identification of a new catalytic motif of oligosaccharyltransferase." Embo Journal 27(1): 234-243.

Irving, G. C. J. and D. J. Cosgrove (1972). "Inositol Phosphate Phosphatases of Microbiological Origin - Inositol Pentaphosphate Products of Aspergillus-Ficuum Phytases." Journal of Bacteriology 112(1): 434-&.

Ito, T., N. Roongsawang, et al. (2009). "Deciphering Pactamycin Biosynthesis and Engineered Production of New Pactamycin Analogues." Chembiochem 10(13): 2253-2265.

Iwasa, T., E. Higashide, et al. (1971). "Studies on validamycins, new antibiotics. II. Production and biological properties of validamycins A and B." The Journal of Antibiotics (Tokyo) 24(2): 107-113.

Iwasa, T., H. Yamamoto, et al. (1970). "Studies on validamycins, new antibiotics. I. Streptomyces hygroscopicus var. limoneus nov. var., validamycin-producing organism." The Japanese Journal of Antibiotics 23(6): 595-602.

Jaenicke, L. (2007). "Centenary of the award of a Nobel prize to Eduard Buchner, the father of biochemistry in a test tube and thus of experimental molecular bioscience." Angewandte Chemie-International Edition 46(36): 6776-6782.

Jamaluddin, H., P. Tumbale, et al. (2007). "Conformational changes induced by binding UDP- 2F-galactose to alpha-1,3-galactosyltransferase - Implications for catalysis." Journal of Molecular Biology 369(5): 1270-1281.

Jinek, M., Y. W. Chen, et al. (2006). "Structural insights into the Notch-modifying glycosyltransferase Fringe." Nature Structural & Molecular Biology 13(10): 945-946.

Kabsch, W. (2010). "Xds." Acta Crystallographica Section D-Biological Crystallography 66: 125-132.

Kakuda, S., T. Shiba, et al. (2004). "Structural basis for acceptor substrate recognition of a human glucuronyltransferase, GlcAT-P, an enzyme critical in the biosynthesis of the carbohydrate epitope HNK-1." Journal of Biological Chemistry 279(21): 22693-22703.

Kameda, Y., N. Asano, et al. (1987). "Validoxylamines as Trehalase Inhibitors." Journal of Antibiotics 40(4): 563-565.

114

Katz, L. (1997). "Manipulation of Modular Polyketide Synthases." Chemical Reviews 97(7): 2557-2576.

Keegan, R. M. and M. D. Winn (2007). "Automated search-model discovery and preparation for structure solution by molecular replacement." Acta Crystallographica Section D- Biological Crystallography 63: 447-457.

Keller, T. H., A. Pichota, et al. (2006). "A practical view of 'druggability'." Current Opinion in Chemical Biology 10(4): 357-361.

Kellogg, B. A. and C. D. Poulter (1997). "Chain elongation in the isoprenoid biosynthetic pathway." Current Opinion in Chemical Biology 1(4): 570-578.

Kennedy, E. J., L. Pillus, et al. (2005). "Pho5p and newly identified nucleotide pyrophosphatases/phosphodiesterases regulate extracellular nucleotide phosphate metabolism in Saccharomyces cerevisiae." Eukaryotic Cell 4(11): 1892-1901.

Kikuchi, N. and H. Narimatsu (2006). "Bioinformatics for comprehensive finding and analysis of glycosyltransferases." Biochimica et Biophysica Acta-General Subjects 1760(4): 578- 583.

Kim, J. W., I. Tchernyshyov, et al. (2006). "HIF-1-mediated expression of pyruvate dehydrogenase kinase: A metabolic switch required for cellular adaptation to hypoxia." Cell Metabolism 3(3): 177-185.

Kim, S. G., M. Cavalier, et al. (2007). "A direct substrate-substrate interaction found in the kinase domain of the bifunctional enzyme, 6-phosphofructo-2-kinase/fructose-2,6- bisphosphatase." Journal of Molecular Biology 370(1): 14-26.

Kim, S. G., N. P. Manes, et al. (2006). "Crystal structure of the hypoxia-inducible form of 6- phosphofructo-2-kinase/fructose-2,6-bisphosphatase (PFKFB3): a possible new target for cancer therapy." Journal of Biological Chemistry 281(5): 2939-2944.

Kingston, W. (2004). "Streptomycin, Schatz v. Waksman, and the balance of credit for discovery." Journal of the History of Medicine and Allied Sciences 59(3): 441-462.

Kinne-Saffran, E. and R. K. Kinne (1999). "Vitalism and synthesis of urea. From Friedrich Wohler to Hans A. Krebs." American Journal of Nephrology 19(2): 290-294.

Kitajima, S., R. Sakakibara, et al. (1984). "Kinetic-Studies of Fructose 6-Phosphate,2-Kinase and Fructose 2,6-Bisphosphatase." Journal of Biological Chemistry 259(11): 6896-6903.

Klahn, M., E. Rosta, et al. (2006). "On the mechanism of hydrolysis of phosphate monoesters dianions in solutions and proteins." Journal of the American Chemical Society 128(47): 15310-15323.

115

Knolker, H. J. and K. R. Reddy (2002). "Isolation and synthesis of biologically active carbazole alkaloids." Chemical Reviews 102(11): 4303-4427.

Koonin, E. V., Y. I. Wolf, et al. (2000). "Protein fold recognition using sequence profiles and its application in structural genomics." Advances in Protein Chemistry 54: 245-275.

Kornberg, S., A. Kornberg, et al. (1961). "Glucosylation of Deoxyribonucleic Acid by Enzymes from Bacteriophage-Infected Escherichia Coli." Journal of Biological Chemistry 236(5): 1487-&.

Kountz, P. D., S. Freeman, et al. (1988). "The stereochemical course of phospho group transfer catalyzed by rat liver 6-phosphofructo-2-kinase." Journal of Biological Chemistry 263(31): 16069-16072.

Krell, T., J. R. Coggins, et al. (1998). "The three-dimensional structure of shikimate kinase." Journal of Molecular Biology 278(5): 983-997.

Kudo, F. and T. Eguchi (2009). "Biosynthetic genes for aminoglycoside antibiotics." Journal of Antibiotics 62(9): 471-481.

Kudo, F., Y. Kasama, et al. (2007). "Cloning of the pactamycin biosynthetic gene cluster and characterization of a crucial glycosyltransferase prior to a unique cyclopentane ring formation." The Journal of Antibiotics (Tokyo) 60(8): 492-503.

Kurland, I. J., B. Chapman, et al. (2000). "N- and C-termini modulate the effects of pH and phosphorylation on hepatic 6-phosphofructo-2-kinase/fructose-2,6-bisphosphatase." Biochemical Journal 347(Pt 2): 459-467.

Kurland, I. J., L. Li, et al. (1993). "Regulation of rat 6-phosphofructo-2-kinase/fructose-2,6- bisphosphatase. Role of the NH2-terminal region." Journal of Biological Chemistry 268(19): 14056-14064.

Lahiri, S. D., G. F. Zhang, et al. (2003). "The pentacovalent phosphorus intermediate of a phosphoryl transfer reaction." Science 299(5615): 2067-2071.

Lairson, L. L., B. Henrissat, et al. (2008). "Glycosyltransferases: Structures, functions, and mechanisms." Annual Review of Biochemistry 77: 521-555.

Langer, G., S. X. Cohen, et al. (2008). "Automated macromolecular model building for X-ray crystallography using ARP/wARP version 7." Nature Protocols 3(7): 1171-1179.

Lariviere, L. and S. Morera (2002). "A base-flipping mechanism for the T4 phage beta- glucosyltransferase and identification of a transition-state analog." Journal of Molecular Biology 324(3): 483-490.

116

Larkin, M. A., G. Blackshields, et al. (2007). "Clustal W and Clustal X version 2.0." Bioinformatics 23(21): 2947-2948.

Lee, S. S., S. Y. Hong, et al. (2011). "Mechanistic evidence for a front-side, S(N)i-type reaction in a retaining glycosyltransferase." Nature Chemical Biology 7(9): 631-638.

Lee, Y. H., Y. Li, et al. (2003). "Tissue-specific structure/function differentiation of the liver isoform of 6-phosphofructo-2-kinase/fructose-2,6-bisphosphatase." Journal of Biological Chemistry 278(1): 523-530.

Lee, Y. H., T. W. Olson, et al. (1997). "Crystal structure of a trapped phosphoenzyme during a catalytic reaction." Nature Structural Biology 4(8): 615-618.

Leipe, D. D., E. V. Koonin, et al. (2003). "Evolution and classification of P-loop kinases and related proteins." Journal of Molecular Biology 333(4): 781-815.

Leipe, D. D., Y. I. Wolf, et al. (2002). "Classification and evolution of P-loop GTPases and related ATPases." Journal of Molecular Biology 317(1): 41-72.

Lin, K., I. J. Kurland, et al. (1994). "Evidence for NH2- and COOH-terminal interactions in rat 6-phosphofructo-2-kinase/fructose-2,6-bisphosphatase." Journal of Biological Chemistry 269(24): 16953-16960.

Lipinski, C. A. (2004). "Lead- and drug-like compounds: the rule-of-five revolution." Drug Discovery Today: Technologies 1(4): 337-341.

Liu, J. and A. Mushegian (2003). "Three monophyletic superfamilies account for the majority of the known glycosyltransferases." Protein Science 12(7): 1418-1431.

Liu, L., A. Thamchaipenet, et al. (1997). "Biosynthesis of 2-nor-6-deoxyerythronolide B by rationally designed domain substitution." Journal of the American Chemical Society 119(43): 10553-10554.

Lizak, C., S. Gerber, et al. (2011). "X-ray structure of a bacterial oligosaccharyltransferase." Nature 474(7351): 350-U377.

Llewellyn, N. M. and J. B. Spencer (2006). "Biosynthesis of 2-deoxystreptamine-containing aminoglycoside antibiotics." Natural Product Reports 23(6): 864-874.

Loiseau, A. M., M. H. Rider, et al. (1988). "Rat Hepatoma (Htc) Cell 6-Phosphofructo-2-Kinase Differs from That in Liver and Can Be Separated from Fructose-2,6-Bisphosphatase." European Journal of Biochemistry 175(1): 27-32.

Lovering, A. L., L. H. de Castro, et al. (2007). "Structural insight into the transglycosylation step of bacterial cell-wall biosynthesis." Science 315(5817): 1402-1405.

117

Lynch, S. R., R. L. Gonzalez, et al. (2003). "Comparison of x-ray crystal structure of the 30S subunit-antibiotic complex with NMR structure of decoding site oligonucleotide- paromomycin complex." Structure 11(1): 43-53.

Macarron, R. (2006). "Critical review of the role of HTS in drug discovery." Drug Discovery Today 11(7–8): 277-279.

Mahmud, T. (2003). "The C7N aminocyclitol family of natural products." Natural Product Reports 20(1): 137-166.

Mahmud, T. (2009). "Progress in aminocyclitol biosynthesis." Current Opinion in Chemical Biology 13(2): 161-170.

Mahmud, T., P. M. Flatt, et al. (2007). "Biosynthesis of unusual aminocyclitol-containing natural products." Journal of Natural Products 70(8): 1384-1391.

Mahmud, T., S. Lee, et al. (2001). "The biosynthesis of acarbose and validamycin." Chemical Record 1(4): 300-310.

Manes, N. P. and M. R. El-Maghrabi (2005). "The kinase activity of human brain 6- phosphofructo-2-kinase/fructose-2,6-bisphosphatase is regulated via inhibition by phosphoenolpyruvate." Archives of Biochemistry and Biophysics 438(2): 125-136.

Mann, R. L., R. M. Gale, et al. (1953). "Hygromycin .2. Isolation and Properties." Antibiotics and Chemotherapy 3(12): 1279-1282.

Manzano, A., J. L. Rosa, et al. (1998). "Molecular cloning, expression, and chromosomal localization of a ubiquitously expressed human 6-phosphofructo-2-kinase/ fructose-2, 6- bisphosphatase gene (PFKFB3)." Cytogenetics and Cell Genetics 83(3-4): 214-217.

Marsin, A. S., L. Bertrand, et al. (2000). "Phosphorylation and activation of heart PFK-2 by AMPK has a role in the stimulation of glycolysis during ischaemia." Current Biology 10(20): 1247-1255.

Marsin, A. S., C. Bouzin, et al. (2002). "The stimulation of glycolysis by hypoxia in activated monocytes is mediated by AMP-activated protein kinase and inducible 6-phosphofructo- 2-kinase." Journal of Biological Chemistry 277(34): 30778-30783.

Martinez-Fleites, C., M. Proctor, et al. (2006). "Insights into the synthesis of lipopolysaccharide and antibiotics through the structures of two retaining glycosyltransferases from family GT4." Chemistry & Biology 13(11): 1143-1152.

Masson, G., C. Philouze, et al. (2005). "Cis-stereoselective SmI2-promoted reductive coupling of keto-nitrones: first synthesis of 1-epitrehazolamine." Organic & Biomolecular Chemistry 3(11): 2067-2069.

118

Maxwell, P. H., C. W. Pugh, et al. (2001). "Activation of the HIF pathway in cancer." Current Opinion in Genetics & Development 11(3): 293-299.

Mccoy, A. J., R. W. Grosse-Kunstleve, et al. (2007). "Phaser crystallographic software." Journal of Applied Crystallography 40: 658-674.

Mcdaniel, R., S. Ebertkhosla, et al. (1995). "Rational Design of Aromatic Polyketide Natural- Products by Recombinant Assembly of Enzymatic Subunits." Nature 375(6532): 549- 554.

McDaniel, R., A. Thamchaipenet, et al. (1999). "Multiple genetic modifications of the erythromycin polyketide synthase to produce a library of novel "unnatural" natural products." Proceedings of the National Academy of Sciences of the United States of America 96(10): 5890-5890.

Mcguire, J. M., R. L. Bunch, et al. (1952). "Ilotycin, a New Antibiotic." Antibiotics and Chemotherapy 2(6): 281-283.

McKee, T. and J. R. McKee (2009). Biochemistry : the molecular basis of life. New York, Oxford University Press.

Merriam-Webster Inc. (2005). The Merriam-Webster dictionary. Springfield, Mass., Merriam- Webster.

Michels, P. A. and D. J. Rigden (2006). "Evolutionary analysis of fructose 2,6-bisphosphate metabolism." IUBMB Life 58(3): 133-141.

Miley, M. J., A. K. Zielinska, et al. (2007). "Crystal structure of the cofactor-binding domain of the human phase II drug-metabolism enzyme UDP-glucuronosyltransferase 2B7." Journal of Molecular Biology 369(2): 498-511.

Minagawa, K., Y. R. Zhang, et al. (2007). "ValC, a new type of C7-cyclitol kinase involved in the biosynthesis of the antifungal agent validamycin A." Chembiochem 8(6): 632-641.

Minchenko, A., I. Leshchinsky, et al. (2002). "Hypoxia-inducible factor-1-mediated expression of the 6-phosphofructo-2-kinase/fructose-2,6-bisphosphatase-3 (PFKFB3) gene. Its possible role in the Warburg effect." Journal of Biological Chemistry 277(8): 6183-6187.

Minchenko, O., I. Opentanova, et al. (2003). "Hypoxic regulation of the 6-phosphofructo-2- kinase/fructose-2,6-bisphosphatase gene family (PFKFB-1-4) expression in vivo." Febs Letters 554(3): 264-270.

Monegal, A. and A. Planas (2006). "Chemical rescue of alpha 3-galactosyltransferase. Implications in the mechanism of retaining glycosyltransferases." Journal of the American Chemical Society 128(50): 16030-16031.

119

Moreno-Sanchez, R., S. Rodriguez-Enriquez, et al. (2007). "Energy metabolism in tumor cells." Febs Journal 274(6): 1393-1418.

Mulichak, A. M., H. C. Losey, et al. (2003). "Structure of the TDP-epi-vancosaminyltransferase GtfA from the chloroeremomycin biosynthetic pathway." Proceedings of the National Academy of Sciences of the United States of America 100(16): 9238-9243.

Mulichak, A. M., H. C. Losey, et al. (2001). "Structure of the UDP-Glucosyltransferase GtfB that modifies the heptapeptide aglycone in the biosynthesis of vancomycin group antibiotics." Structure 9(7): 547-557.

Mulichak, A. M., W. Lu, et al. (2004). "Crystal structure of vancosaminyltransferase GtfD from the vancomycin biosynthetic pathway: Interactions with acceptor and nucleotide ligands." Biochemistry 43(18): 5170-5180.

Muller, C. W. and G. E. Schulz (1992). "Structure of the Complex between Adenylate Kinase from Escherichia-Coli and the Inhibitor Ap5a Refined at 1.9 a Resolution - a Model for a Catalytic Transition-State." Journal of Molecular Biology 224(1): 159-177.

Murray, K. J., M. R. Elmaghrabi, et al. (1984). "Amino-Acid-Sequence of the Phosphorylation Site of Rat-Liver 6-Phosphofructo-2-Kinase Fructose-2,6-Bisphosphatase." Journal of Biological Chemistry 259(12): 7673-7681.

Nango, E., T. Kumasaka, et al. (2008). "Structure of 2-deoxy-scyllo-inosose synthase, a key enzyme in the biosynthesis of 2-deoxystreptamine-containing aminoglycoside antibiotics, in complex with a mechanism-based inhibitor and NAD+." Proteins-Structure Function and Bioinformatics 70(2): 517-527.

Newman, D. J. and G. M. Cragg (2012). "Natural Products As Sources of New Drugs over the 30 Years from 1981 to 2010." Journal of Natural Products 75(3): 311-335.

O'Hagan, D. (1991). The Polyketide Metabolites. Chichester, U. K., Horwood.

Obach, M., A. Navarro-Sabate, et al. (2004). "6-Phosphofructo-2-kinase (pfkfb3) gene promoter contains hypoxia-inducible factor-1 binding sites necessary for transactivation in response to hypoxia." Journal of Biological Chemistry 279(51): 53562-53570.

Offen, W., C. Martinez-Fleites, et al. (2006). "Structure of a flavonoid glucosyltransferase reveals the basis for plant natural product modification." Embo Journal 25(6): 1396-1405.

Ogle, J. M. and V. Ramakrishnan (2005). "Structural insights into translational fidelity." Annual Review of Biochemistry 74: 129-177.

Okamura, N. and R. Sakakibara (1998). "A common phosphorylation site for cyclic AMP- dependent protein kinase and protein kinase C in human placental 6-phosphofructo-2-

120

kinase/fructose-2,6-bisphosphatase." Bioscience Biotechnology and Biochemistry 62(10): 2039-2042.

Olesen, C., T. L. Sorensen, et al. (2004). "Dephosphorylation of the calcium pump coupled to counterion occlusion." Science 306(5705): 2251-2255.

Oliynyk, M., M. J. B. Brown, et al. (1996). "A hybrid modular polyketide synthase obtained by domain swapping." Chemistry & Biology 3(10): 833-839.

Omura, S., A. Nakagawa, et al. (1987). "Hygromycin-a, an Antitreponemal Substance .1. Screening Method and Therapeutic Effect for Treponema-Hyodysenteriae-Caused Infection in Cf-1 Mice." Journal of Antibiotics 40(11): 1619-1626.

Onaka, H., S. Taniguchi, et al. (2002). "Cloning of the staurosporine biosynthetic gene cluster from Streptomyces sp TP-A0274 and its heterologous expression in Streptomyces lividans." Journal of Antibiotics 55(12): 1063-1071.

Onaka, H., S. Taniguchi, et al. (2003). "Characterization of the biosynthetic gene cluster of rebeccamycin from Lechevalieria aerocolonigenes ATCC 39243." Bioscience Biotechnology and Biochemistry 67(1): 127-138.

Otwinowski, Z. and W. Minor (1997). "Processing of X-ray diffraction data collected in oscillation mode." Macromolecular Crystallography, Pt A 276: 307-326.

Painter, J. and E. A. Merritt (2006). "Optimal description of a protein structure in terms of multiple groups undergoing TLS motion." Acta Crystallographica Section D-Biological Crystallography 62: 439-450.

Painter, J. and E. A. Merritt (2006). "TLSMD web server for the generation of multi-group TLS models." Journal of Applied Crystallography 39: 109-111.

Pak, J. E., P. Arnoux, et al. (2006). "X-ray crystal structure of leukocyte type core 2 beta 1,6-N- acetylglucosaminyltransferase - Evidence for a convergence of metal ion-independent glycosyltransferase mechanism." Journal of Biological Chemistry 281(36): 26693-26701.

Pal, S. (2006). "A journey across the sequential development of macrolides and ketolides related to erythromycin." Tetrahedron 62(14): 3171-3200.

Papandreou, I., R. A. Cairns, et al. (2006). "HIF-1 mediates adaptation to hypoxia by actively downregulating mitochondrial oxygen consumption." Cell Metabolism 3(3): 187-197.

Parsonage, D., G. L. Newton, et al. (2010). "Characterization of the N-Acetyl-alpha-D- glucosaminyl L-Malate Synthase and Deacetylase Functions for Bacillithiol Biosynthesis in Bacillus anthracis." Biochemistry 49(38): 8398-8414.

121

Pedersen, L. C., K. Tsuchida, et al. (2000). "Heparan/chondroitin sulfate biosynthesis - Structure and mechanism of human glucuronyltransferase I." Journal of Biological Chemistry 275(44): 34580-34585.

Pelicano, H., D. S. Martin, et al. (2006). "Glycolysis inhibition for anticancer treatment." Oncogene 25(34): 4633-4646.

Persson, K., H. D. Ly, et al. (2001). "Crystal structure of the retaining galactosyltransferase LgtC from Neisseria meningitidis in complex with donor and acceptor sugar analogs." Nature Structural Biology 8(2): 166-175.

Pfeiffer, T., S. Schuster, et al. (2001). "Cooperation and competition in the evolution of ATP- producing pathways." Science 292(5516): 504-507.

Pilkis, S. J., T. Chrisman, et al. (1983). "Rat Hepatic 6-Phosphofructo 2-Kinase Fructose 2,6- Bisphosphatase - a Unique Bifunctional Enzyme." Advances in Enzyme Regulation 21: 147-&.

Pilkis, S. J., M. R. El-Maghrabi, et al. (1981). "Inhibition of fructose-1,6-bisphosphatase by fructose 2,6-bisphosphate." Journal of Biological Chemistry 256(8): 3619-3622.

Pilkis, S. J., M. Walderhaug, et al. (1983). "6-Phosphofructo-2-Kinase Fructose 2,6- Bisphosphatase from Rat-Liver - Isolation and Identification of a Phosphorylated Intermediate." Journal of Biological Chemistry 258(10): 6135-6141.

Pittenger, R. C., R. N. Wolfe, et al. (1953). "Hygromycin .1. Preliminary Studies on the Production and Biologic Activity of a New Antibiotic." Antibiotics and Chemotherapy 3(12): 1268-1278.

Potterton, E., P. Briggs, et al. (2003). "A graphical user interface to the CCP4 program suite." Acta Crystallographica Section D-Biological Crystallography 59: 1131-1137.

Prasad, R., S. K. Verma, et al. (2006). "Efficacy and safety of kanamycin, ethionamide, PAS and cycloserine in multidrug-resistant pulmonary tuberculosis patients." The Indian Journal of Chest Diseases & Allied Sciences 48(3): 183-186.

Prudhomme, M. (2003). "Rebeccamycin Analogues as Anticancer Agents." ChemInform 34(23): no-no.

Puls, W., U. Keup, et al. (1977). "Glucosidase Inhibition - New Approach to Treatment of Diabetes, Obesity, and Hyperlipoproteinemia." Naturwissenschaften 64(10): 536-537.

Punta, M., P. C. Coggill, et al. (2012). "The Pfam protein families database." Nucleic Acids Research 40(D1): D290-D301.

122

Ramakrishnan, B., V. Ramasamy, et al. (2006). "Structural snapshots of beta-1,4- galactosyltransferase-1 along the kinetic pathway." Journal of Molecular Biology 357(5): 1619-1633.

Rebora, K., B. Laloo, et al. (2005). "Revisiting purine-histidine cross-pathway regulation in Saccharomyces cerevisiae: A central role for a small molecule." Genetics 170(1): 61-70.

Reinhart, G. D. and H. A. Lardy (1980). "Rat-Liver Phosphofructokinase - Kinetic Activity under near-Physiological Conditions." Biochemistry 19(7): 1477-1484.

Richards, C. S. and K. Uyeda (1980). "Changes in the Concentration of Activation Factor for Phosphofructokinase in Hepatocytes in Response to Glucose and Glucagon." Biochemical and Biophysical Research Communications 97(4): 1535-1540.

Richards, C. S., M. Yokoyama, et al. (1982). "Reciprocal Changes in Fructose-6-Phosphate, 2- Kinase and Fructose-2,6-Bisphosphatase Activity in Response to Glucagon and Epinephrine." Biochemical and Biophysical Research Communications 104(3): 1073- 1079.

Rider, M. H., D. Foret, et al. (1985). "Comparison of purified bovine heart and rat liver 6- phosphofructo-2-kinase. Evidence for distinct isoenzymes." Biochemical Journal 231(1): 193-196.

Rigden, D. J. (2008). "The histidine phosphatase superfamily: structure and function." Biochemical Journal 409: 333-348.

Rinehart, K. L., D. D. Weller, et al. (1980). "Recent Biosynthetic-Studies on Antibiotics." Journal of Natural Products 43(1): 1-20.

Rittinger, K., P. A. Walker, et al. (1997). "Structure at 1.65 A of RhoA and its GTPase-activating protein in complex with a transition-state analogue." Nature 389(6652): 758-762.

Rose, Z. B. (1970). "Evidence for a Phosphohistidine Protein Intermediate in Phosphoglycerate Mutase Reaction." Archives of Biochemistry and Biophysics 140(2): 508-&.

Ruan, X., A. Pereda, et al. (1997). "Acyltransferase domain substitutions in erythromycin polyketide synthase yield novel erythromycin derivatives." Journal of Bacteriology 179(20): 6416-6425.

Sakai, A., M. Kato, et al. (1996). "Cloning of cDNA encoding for a novel isozyme of fructose 6- phosphate, 2-kinase/fructose 2,6-bisphosphatase from human placenta." Journal of Biochemistry 119(3): 506-511.

Sakakibara, R., M. Kato, et al. (1997). "Characterization of a human placental fructose-6- phosphate 2-kinase fructose-2,6-bisphosphatase." Journal of Biochemistry 122(1): 122- 128.

123

Sakakibara, R., S. Kitajima, et al. (1984). "Differences in Kinetic-Properties of Phospho and Dephospho Forms of Fructose-6-Phosphate,2-Kinase and Fructose 2,6-Bisphosphatase." Journal of Biological Chemistry 259(1): 41-46.

Sakakibara, R., T. Okudaira, et al. (1999). "Tissue distribution of placenta-type 6-phosphofructo- 2-kinase/fructose-2,6-bisphosphatase." Biochemical and Biophysical Research Communications 257(1): 177-181.

Sakata, J., Y. Abe, et al. (1991). "Molecular cloning of the DNA and expression and characterization of rat testes fructose-6-phosphate,2-kinase:fructose-2,6-bisphosphatase." Journal of Biological Chemistry 266(24): 15764-15770.

Sakuda, S., A. Isogai, et al. (1986). "The Structure of Allosamidin, a Novel Insect Chitinase Inhibitor, Produced by Streptomyces Sp." Tetrahedron Letters 27(22): 2475-2478.

Sakuda, S., Y. Sugiyama, et al. (2001). "Biosynthetic studies on the cyclopentane ring formation of allosamizoline, an aminocyclitol component of the chitinase inhibitor allosamidin." Journal of Organic Chemistry 66(10): 3356-3361.

Sakurai, M., C. A. Hasemann, et al. (2000). "Activation of Glu325AIa mutant enzyme of fructose 6-phosphate 2-kinase: Fructose 2,6-bisphosphatase by exogenous nucleophiles." Faseb Journal 14(8): A1523-A1523.

Salceda, S. and J. Caro (1997). "Hypoxia-inducible factor 1 alpha (HIF-1 alpha) protein is rapidly degraded by the ubiquitin-proteasome system under normoxic conditions - Its stabilization by hypoxia depends on redox-induced changes." Journal of Biological Chemistry 272(36): 22642-22647.

Sanchez, C., C. Mendez, et al. (2006). "Engineering biosynthetic pathways to generate antitumor indolocarbazole derivatives." Journal of Industrial Microbiology & Biotechnology 33(7): 560-568.

Sanchez, C., L. L. Zhu, et al. (2005). "Combinatorial biosynthesis of antitumor indolocarbazole compounds." Proceedings of the National Academy of Sciences of the United States of America 102(2): 461-466.

Schlorke, O., P. Krastel, et al. (2002). "Structure and biosynthesis of cetoniacytone A, a cytotoxic aminocarba sugar produced by an endosymbiontic Actinoimyces." Journal of Antibiotics 55(7): 635-642.

Schmidt, D. D., W. Frommer, et al. (1977). "Alpha-Glucosidase Inhibitors - New Complex Oligosaccharides of Microbial Origin." Naturwissenschaften 64(10): 535-536.

Schneider, G., Y. Lindqvist, et al. (1993). "3-Dimensional Structure of Rat Acid-Phosphatase." Embo Journal 12(7): 2609-2615.

124

Semenza, G. L. (2002). "HIF-1 and tumor progression: pathophysiology and therapeutics." Trends in Molecular Medicine 8(4): S62-S67.

Semenza, G. L. (2006). "Development of novel therapeutic strategies that target HIF-1." Expert Opinion on Therapeutic Targets 10(2): 267-280.

Seo, M., J. D. Kim, et al. (2011). "Structure-Based Development of Small Molecule PFKFB3 Inhibitors: A Framework for Potential Cancer Therapeutic Agents Targeting the Warburg Effect." PLoS One 6(9).

Seo, M. J., E. M. Im, et al. (2006). "Characterization of D-glucose alpha-1-phosphate uridylyltransferase (VIdB) and glucokinase (VldC) involved in validamycin biosynthesis of Streptomyces hygroscopicus var. limoneus KCCM 11405." Journal of Microbiology and Biotechnology 16(8): 1311-1315.

Shao, H., X. Z. He, et al. (2005). "Crystal structures of a multifunctional triterpene/flavonoid glycosyltransferase from Medicago truncatula." Plant Cell 17(11): 3141-3154.

Shibata, M., M. Uyeda, et al. (1981). "Stimulation of the extension of validamycin-inhibited hyphae by the hyphal extension factor present in Rhizoctonia solani." The Journal of Antibiotics (Tokyo) 34(4): 447-451.

Shick, J. M. and W. C. Dunlap (2002). "Mycosporine-like amino acids and related gadusols: Biosynthesis, accumulation, and UV-protective functions in aquatic organisms." Annual Review of Physiology 64: 223-262.

Shigemoto, R., T. Okuno, et al. (1992). "Effects of Validamycin A on the Growth of and Trehalose Content in Mycelia of Rhizoctonia solani Incubated in a Medium Containing Several Sugars as the Sole Carbon Source." Japanese Journal of Phytopathology 58(5): 685-690.

Shim, H., Y. S. Chun, et al. (1998). "A unique glucose-dependent apoptotic pathway induced by c-Myc." Proceedings of the National Academy of Sciences of the United States of America 95(4): 1511-1516.

Singh, D., M. J. Seo, et al. (2006). "Genetic localization and heterologous expression of validamycin biosynthetic gene cluster isolated from Streptomyces hygroscopicus var. limoneus KCCM 11405 (IFO 12704)." Gene 376(1): 13-23.

Stassi, D. L., S. J. Kakavas, et al. (1998). "Ethyl-substituted erythromycin derivatives produced by directed metabolic engineering." Proceedings of the National Academy of Sciences of the United States of America 95(13): 7305-7309.

Stragier, P. and R. Losick (1996). "Molecular genetics of sporulation in Bacillus subtilis." Annual Review of Genetics 30: 297-341.

125

Sugiyama, Y., H. Nagasawa, et al. (2002). "Biosynthesis of the trehalase inhibitor trehazolin." The Journal of Antibiotics (Tokyo) 55(3): 263-269.

Telang, S., A. Yalcin, et al. (2006). "Ras transformation requires metabolic control by 6- phosphofructo-2-kinase." Oncogene 25(55): 7225-7234.

Terwilliger, T. C. (2004). "Using prime-and-switch phasing to reduce model bias in molecular replacement." Acta Crystallographica Section D-Biological Crystallography 60: 2144- 2149.

Terwilliger, T. C., R. W. Grosse-Kunstleve, et al. (2008). "Iterative-build OMIT maps: map improvement by iterative model building and refinement without model bias." Acta Crystallographica Section D-Biological Crystallography 64: 515-524.

Terwilliger, T. C., R. W. Grosse-Kunstleve, et al. (2008). "Iterative model building, structure refinement and density modification with the PHENIX AutoBuild wizard." Acta Crystallographica Section D-Biological Crystallography 64: 61-69.

Tews, I., A. Perrakis, et al. (1996). "Bacterial chitobiase structure provides insight into catalytic mechanism and the basis of Tay-Sachs disease." Nature Structural Biology 3(7): 638-648.

Thornton, R. D., P. Lane, et al. (2000). "Interleukin 1 induces hypoxia-inducible factor 1 in human gingival and synovial fibroblasts." Biochemical Journal 350: 307-312.

Thulasiram, H. V. and C. D. Poulter (2006). "Farnesyl diphosphate synthase: The art of compromise between substrate selectivity and stereoselectivity." Journal of the American Chemical Society 128(49): 15819-15823.

Traut, T. W. (1994). "The Functions and Consensus Motifs of 9 Types of Peptide Segments That Form Different Types of Nucleotide-Binding Sites." European Journal of Biochemistry 222(1): 9-19.

Truscheit, E., W. Frommer, et al. (1981). "Chemistry and Biochemistry of Microbial Alpha- Glucosidase Inhibitors." Angewandte Chemie-International Edition 20(9): 744-761.

Tvaroska, I. (2006). "Molecular modeling of retaining glycosyltransferases." NMR Spectroscopy and Computer Modeling of Carbohydrates: Recent Advances 930: 285-301.

Unligil, U. M., S. H. Zhou, et al. (2000). "X-ray crystal structure of rabbit N- acetylglucosaminyltransferase I: catalytic mechanism and a new protein superfamily." Embo Journal 19(20): 5269-5280.

Uyeda, K., E. Furuya, et al. (1982). "Fructose-2,6-P2, Chemistry and Biological Function." Molecular and Cellular Biochemistry 48(2): 97-120.

126

Uyeda, K., E. Furuya, et al. (1981). "The Structure of Activation Factor for Phosphofructokinase." Journal of Biological Chemistry 256(16): 8679-8684.

Vagin, A. and A. Teplyakov (1997). "MOLREP: an automated program for molecular replacement." Journal of Applied Crystallography 30: 1022-1025.

Vagin, A. A., R. A. Steiner, et al. (2004). "REFMAC5 dictionary: organization of prior chemical knowledge and guidelines for its use." Acta Crystallographica Section D-Biological Crystallography 60: 2184-2195.

Van Schaftingen, E., D. R. Davies, et al. (1981). "Inactivation of Phosphofructokinase-2 by Cyclic Amp-Dependent Protein-Kinase." Biochemical and Biophysical Research Communications 103(1): 362-368.

Van Schaftingen, E., D. R. Davies, et al. (1982). "Fructose-2,6-Bisphosphatase from Rat-Liver." European Journal of Biochemistry 124(1): 143-149.

Van Schaftingen, E. and H. G. Hers (1980). "Synthesis of a Stimulator of Phosphofructokinase, Most Likely Fructose 2,6-Bis-Phosphate, from Phosphoric-Acid and Fructose 6- Phosphoric Acid." Biochemical and Biophysical Research Communications 96(4): 1524- 1531.

Van Schaftingen, E., L. Hue, et al. (1980). "Control of the Fructose 6-Phosphate-Fructose 1,6- Bisphosphate Cycle in Isolated Hepatocytes by Glucose and Glucagon - Role of a Low Molecular-Weight Stimulator of Phosphofructokinase." Biochemical Journal 192(3): 887-895.

Van Schaftingen, E., L. Hue, et al. (1980). "Fructose 2,6-Bisphosphate, the Probable Structure of the Glucose- and Glucagon-Sensitive Stimulator of Phosphofructokinase." Biochemical Journal 192(3): 897-901.

Vanetten, R. L. and M. E. Hickey (1977). "Phosphohistidine as a Stoichiometric Intermediate in Reactions Catalyzed by Isoenzymes of Wheat-Germ Acid-Phosphatase." Archives of Biochemistry and Biophysics 183(1): 250-259.

Ventura, F., J. L. Rosa, et al. (1992). "Bovine Brain 6-Phosphofructo-2-Kinase Fructose-2,6- Bisphosphatase - Evidence for a Neural-Specific Isozyme." Journal of Biological Chemistry 267(25): 17939-17943.

Vertesy, L., H. W. Fehlhaber, et al. (1994). "The Trehalase Inhibitor Salbostatin, a Novel Metabolite from Streptomyces-Albus, Atcc21838." Angewandte Chemie-International Edition 33(18): 1844-1846.

Vertommen, D., L. Bertrand, et al. (1996). "The ATP-binding site in the 2-kinase domain of liver 6-phosphofructo-2-kinase/fructose-2,6-bisphosphatase - Study of the role of Lys-54 and

127

Thr-55 by site-directed mutagenesis." Journal of Biological Chemistry 271(30): 17875- 17880.

Vester, B. and S. Douthwaite (2001). "Macrolide resistance conferred by base substitutions in 23S rRNA." Antimicrob Agents Chemother 45(1): 1-12.

Vetting, M. W., P. A. Frantom, et al. (2008). "Structural and enzymatic analysis of MshA from Corynebacterium glutamicum - Substrate-assisted catalysis." Journal of Biological Chemistry 283(23): 15834-15844.

Vocadlo, D. J., G. J. Davies, et al. (2001). "Catalysis by hen egg-white lysozyme proceeds via a covalent intermediate." Nature 412(6849): 835-838.

Vrielink, A., W. Ruger, et al. (1994). "Crystal structure of the DNA modifying enzyme beta- glucosyltransferase in the presence and absence of the substrate uridine diphosphoglucose." Embo Journal 13(15): 3413-3422.

Walker, J. E., M. Saraste, et al. (1982). "Distantly Related Sequences in the Alpha-Subunits and Beta-Subunits of Atp Synthase, Myosin, Kinases and Other Atp-Requiring Enzymes and a Common Nucleotide Binding Fold." Embo Journal 1(8): 945-951.

Warburg, O. (1956). "On the origin of cancer cells." Science 123(3191): 309-314.

Wechalekar, K., B. Sharma, et al. (2005). "PET/CT in oncology--a major advance." Clinical Radiology 60(11): 1143-1155.

Wehmeier, U. F. and W. Piepersberg (2004). "Biotechnology and molecular biology of the alpha-glucosidase inhibitor acarbose." Applied Microbiology and Biotechnology 63(6): 613-625.

Weller, D. D. and K. L. Rinehart (1978). "Biosynthesis of Anti-Tumor Antibiotic Pactamycin - Methionine-Derived Ethyl Group and a C7n Unit." Journal of the American Chemical Society 100(21): 6757-6760.

Wiley, P. F., H. K. Jahnke, et al. (1970). "Structure of Pactamycin." Journal of Organic Chemistry 35(5): 1420-&.

Williams, A. C., T. J. Collard, et al. (1999). "An acidic environment leads to p53 dependent induction of apoptosis in human adenoma and carcinoma cell lines: implications for clonal selection during colorectal carcinogenesis." Oncogene 18(21): 3199-3204.

Winchester, B. and G. W. J. Fleet (1992). "Amino-Sugar Glycosidase Inhibitors - Versatile Tools for Glycobiologists." Glycobiology 2(3): 199-210.

Wittinghofer, A. (2006). "Phosphoryl transfer in Ras proteins, conclusive or elusive?" Trends in Biochemical Sciences 31(1): 20-23.

128

Woo, E. J., S. I. Ryu, et al. (2010). "Structural Insights on the New Mechanism of Trehalose Synthesis by Trehalose Synthase TreT from Pyrococcus horikoshii." Journal of Molecular Biology 404(2): 247-259.

Woodward, R. B., E. Logusch, et al. (1981). "Asymmetric Total Synthesis of Erythromycin .3. Total Synthesis of Erythromycin." Journal of the American Chemical Society 103(11): 3215-3217.

Wu, X., P. M. Flatt, et al. (2007). "A comparative analysis of the sugar phosphate cyclase superfamily involved in primary and secondary metabolism." Chembiochem 8(2): 239- 248.

Wu, X. M., P. M. Flatt, et al. (2009). "Biosynthetic Gene Cluster of Cetoniacytone A, an Unusual Aminocyclitol from the Endosymbiotic Bacterium Actinomyces sp Lu 9419." Chembiochem 10(2): 304-314.

Xia, T. H. and R. S. Jiao (1986). "Studies on Glutamine-Synthetase from Streptomyces- Hygroscopicus Var Jinggangensis." Scientia Sinica Series B-Chemical Biological Agricultural Medical & Earth Sciences 29(4): 379-388.

Xu, H., K. Minagawa, et al. (2008). "Catalytic analysis of the validamycin glycosyltransferase (ValG) and enzymatic production of 4 ''-epi-validamycin A." Journal of Natural Products 71(7): 1233-1236.

Xu, H., J. T. Yang, et al. (2009). "Genetically engineered production of 1,1'-bis-valienamine and validienamycin in Streptomyces hygroscopicus and their conversion to valienamine." Applied Microbiology and Biotechnology 81(5): 895-902.

Xu, H., Y. R. Zhang, et al. (2009). "Alternative Epimerization in C7N-Aminocyclitol Biosynthesis Is Catalyzed by ValD, A Large Protein of the Vicinal Oxygen Chelate Superfamily." Chemistry & Biology 16(5): 567-576.

Xu, R. H., H. Pelicano, et al. (2005). "Inhibition of glycolysis in cancer cells: A novel strategy to overcome drug resistance associated with mitochondrial respiratory defect and hypoxia." Cancer Research 65(2): 613-621.

Yamamura, S., K. Kawada, et al. (2008). "Prediction of aminoglycoside response against methicillin-resistant Staphylococcus aureus infection in burn patients by artificial neural network modeling." Biomedicine & Pharmacotherapy 62(1): 53-58.

Yang, J. T., H. Xu, et al. (2011). "Nucleotidylation of unsaturated carbasugar in validamycin biosynthesis." Organic & Biomolecular Chemistry 9(2): 438-449.

129

Yang, Y., H. Boze, et al. (2009). "NMR Structure of rALF-Pm3, an Anti-Lipopolysaccharide Factor from Shrimp: Model of the Possible Lipid A-Binding Site." Biopolymers 91(3): 207-220.

Yu, Y., L. Q. Bai, et al. (2005). "Gene cluster responsible for validamycin biosynthesis in Streptomyces hygroscopicus subsp jinggangensis 5008." Applied and Environmental Microbiology 71(9): 5066-5076.

Yuen, M. H., H. Mizuguchi, et al. (1999). "Crystal structure of the H256A mutant of rat testis fructose-6-phosphate,2-kinase/fructose-2,6-bisphosphatase. Fructose 6-phosphate in the active site leads to mechanisms for both mutant and wild type bisphosphatase activities." Journal of Biological Chemistry 274(4): 2176-2184.

Zayas, C. L. and J. C. Escalante-Semerena (2007). "Reassessment of the late steps of coenzyme B-12 synthesis in Salmonella enterica: Evidence that dephosphorylation of adenosylcobalamin-5 '-phosphate by the CobC phosphatase is the last step of the pathway." Journal of Bacteriology 189(6): 2210-2218.

Zhang, C. S., A. Stratmann, et al. (2002). "Biosynthesis of the C(7)-cyclitol moiety of acarbose in Actinoplanes species SE50/110. 7-O-phosphorylation of the initial cyclitol precursor leads to proposal of a new biosynthetic pathway." Journal of Biological Chemistry 277(25): 22853-22862.

Zheng, L., X. Zhou, et al. (2012). "Structural and Functional Analysis of Validoxylamine A 7'- phosphate Synthase ValL Involved in Validamycin A Biosynthesis." PLoS One 7(2): e32033.

Zhou, J. and B. Brune (2006). "Cytokines and hormones in the regulation of hypoxia inducible factor-1alpha (HIF-1alpha)." Cardiovascular and Hematological Agents in Medicinal Chemistry 4(3): 189-197.

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APPENDIXES

A. A Crystallographic Study

X-ray crystallography is a form of very high resolution microscopy that allows researchers to generate a three-dimensional structure of a protein and is one of the few techniques that can provide information regarding the catalytic site at the atomic, or near- atomic, level. This technique is implemented by growing a crystal comprised entirely of protein and then exposing it to an x-ray source. The diffracted x-rays are then measured and used to interpret the position of each atom in three-dimensional space. Once the positions of the protein atoms are calculated, researchers can investigate the interactions between the protein and its bound ligands. This information can explain how ligands influence a protein and which amino acids are involved in their binding. The specific amino acids involved in catalysis can also be identified.

Furthermore, x-ray crystallography can be used to support or explain the observations made in kinetic and functional studies of enzymes.

Within this document, x-ray crystallography was used to investigate the catalysis performed by the enzymes, PFKFB3 and VldE. Using this technique, the intermediates of the PFKFB3 reaction were successfully visualized and the catalytic mechanism responsible for the retention of the stereochemical configuration within VldE products was elucidated. X-ray crystallography allowed a visual interpretation of these two reactions which would be otherwise unattainable.

131

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VITA

Michael Christopher Cavalier is the eldest son of Nickie Cavalier and Bonnie H.

Cavalier. He was born in Baton Rouge, Louisiana on June 17, 1984. He was raised in Pierre

Part, Louisiana. He attended Assumption High School in Napoleonville, Louisiana, where he graduated as co-valedictorian in 2002. He then moved to Baton Rouge, Louisiana, where he received his Bachelor of Science degree in biochemistry from Louisiana State University. He then joined the laboratory of Dr. Yong-Hwan Lee as a graduate assistant in the fall of 2006.

Michael will be defending his dissertation in the summer of 2012 and he will await the issuing of his Doctor of Philosophy degree in Biochemistry in December 2012. His future plans include moving to Baltimore, Maryland, to continue his research in cancer biology as a postdoctoral researcher in the laboratory of Dr. David J. Weber at The University of Maryland, School of

Medicine.

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