A FUNCTIONAL AZASUGAR BIOSYNTHETIC CLUSTER FROM CHITINOPHAGA PINENSIS

By

CLARIBEL NUÑEZ

A DISSERTATION PRESENTED TO THE GRADUATE SCHOOL OF THE UNIVERSITY OF FLORIDA IN PARTIAL FULFILLMENT OF THE REQUIREMENTS FOR THE DEGREE OF DOCTOR OF PHILOSOPHY

UNIVERSITY OF FLORIDA

2019

© 2019 Claribel Nuñez

To me

ACKNOWLEDGMENTS

I am grateful to many people that have supported me throughout my graduate school journey.

Firstly, I would like to thank my mentor, Professor Nicole A. Horenstein for giving me the opportunity to work in her lab. She has been one of my biggest advocates and one of the major forces that kept me striding through both personal issues and graduate school work over the past five years. Much of what I have accomplished and how I have evolved as a scientist is thanks to her guidance and encouragement. I would also like to thank my committee members

Prof. Jon Stewart, Prof. Rebecca Butcher, Prof. Ron Castellano, and Prof. Valerie De Crecy for their advice throughout my research, as well as access to their laboratory equipment and lab space. In addition, I would like to thank Dr. Ion Ghiviriga for his help in analyzing my NMR samples, as well as the Mass Spectrometry facility at UF for their assistance in analyzing my samples, especially Dr. Kari Basso and Dr. Manasi Kamat.

Secondly, thank you to my friends and family for providing their encouragement and understanding. I am very fortunate to have met so many wonderful friends that have supported me as a graduate student. I’m grateful for “El Corillo”—Carla, Rebeca, Johnny, Jose, Rene,

Kathy, Lorena, Paul, Glenda, Angie, Camilo, Christian, Lorraine, Andreina, Gabriel—for our

Latin dance nights, karaoke nights, and cookouts that helped pick me up when I was feeling homesick. Especially my dear friends, Dr. Veronica Negron and Dr. Melissa Cruz-Acuña for feeding me when I was sick and for reassuring me whenever I lost confidence and motivation.

Grateful for my roommate Lindsey for her words of encouragement and for helping me take care of my pup, Oaklee, whenever I had to work late nights in the lab. I’m grateful for my best friend,

Trisha Ramdihal, for our late nights of de-stressing and motivational conversations, that helped me push forward on bad days. I would also like to extend my gratitude to my godfather, Dr.

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Thomas Jordan, for being someone that I could always count on by providing emotional and financial support throughout my years as a graduate student.

Thirdly, I’d like to acknowledge my friends and colleagues in the Chemistry department that have helped me along the way. Thankful for the past and present members of the Horenstein group, including Dr. Jeffrey Arciola, Dr. Alican Gulsevin, Dr. Timothy Gould, Maria Chiara

Pismataro, Hailey Beal, Haoxi Li, and Dr. Marta Quadri. I would like to extend my appreciation to Marta for not only being a wonderful colleague, discussing research results and ideas even though my work was not her expertise, but also for being a great friend; her wonderful meals that helped keep me alive, her ability to help me save money by always finding deals and coupons when grocery shopping, and for sometimes being the only person willing to go out with me to help release some stress even though we were always so busy. I’m grateful for my wonderful friends Dr. Michelle Nolan, Dr. Danielle Fagnani, and Kathryn Olsen for always making mental health checks, for providing scientific advice and always being available to help edit any of my documents. I would not have survived graduate school without the help and support from each and every one of these individuals, and for that I am eternally grateful.

Finally, I thank the University of Florida and the Florida Education Fund’s McKnight

Doctoral Fellowship for their financial support over my doctoral study period.

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TABLE OF CONTENTS

page

ACKNOWLEDGMENTS ...... 4

LIST OF TABLES ...... 8

LIST OF FIGURES ...... 9

LIST OF ABBREVIATIONS...... 13

ABSTRACT ...... 16

CHAPTER

1 INTRODUCTION ...... 19

1.1 Introduction to Secondary Metabolites ...... 19 1.2 Glycosidase Inhibitors ...... 20 1.3 Azasugars ...... 21 1.3.1 Overview ...... 21 1.3.2 Therapeutic Potential and Applications of Azasugars ...... 23 1.3.2.1 Azasugars as antidiabetic agents ...... 23 1.3.2.2 Azasugars in lysosomal storage diseases ...... 23 1.3.2.3 Additional potential therapeutic applications of azasugars ...... 25 1.3.3 Biosynthetic Pathways of Azasugars ...... 26 1.3.3.1 Biosynthesis of nojirimycins ...... 26 1.3.3.2 Biosynthesis of DMPD ...... 29 1.3.3.3 Biosynthesis of nectrisine ...... 29

2 IDENTIFICATION AND CHARACTERIZATION OF DAB-1 IN C. PINENSIS ...... 31

2.1 Overview ...... 31 2.2 Results and Discussion ...... 32 2.3 Experimental ...... 43 2.3.1 General Methods ...... 43 2.3.2 Growth Conditions for Chitinophaga pinensis and Culture Extract Purification ...... 43 2.3.2.1 Rich media growth conditions ...... 43 2.3.2.2 Minimal media growth conditions ...... 44 2.3.2.3 Purification methods ...... 44 2.3.3 Glycosidase Inhibition Assays ...... 47 2.3.3.1  and -glucosidase assays ...... 47 2.3.3.2 Assays on multi-well plate reader ...... 47 2.3.3.3 Jack Bean -mannosidase assay ...... 47

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3 HETEROLOGOUS EXPRESSION OF AZASUGAR BIOSYNTHETIC SIGNATURE IN E. COLI ...... 49

3.1 Overview ...... 49 3.2 Results and Discussion ...... 49 3.3 Experimental ...... 62 3.3.1 General Methods ...... 62 3.3.2 Cloning of the pETBlue2-Cpin2154-Cpin2153-Cpin2152 Expression Construct ...... 62 3.3.3 Expression of Cpin2154, Cpin2153, Cpin2152 in E. coli and purification of nectrisine ...... 63 3.3.3.1 Rich media growth ...... 63 3.3.3.2 Minimal media growth ...... 64 3.3.3.3 Purification methods for compounds produced in E. coli ...... 64 3.3.4 Glycosidase Inhibition Assays ...... 65

4 BIOINFORMATIC ANALYSIS OF THE AZASUGAR BIOSYNTHETIC SIGNATURE ...... 66

4.1 Overview ...... 66 4.2 Results and Discussion ...... 68 4.3 Experimental ...... 83

5 CONCLUSIONS AND FUTURE DIRECTIONS ...... 84

5.1 Conclusions ...... 84 5.2 Future Directions ...... 85

APPENDIX

A GLYCOSIDASE INHIBITION ASSAY FIGURES ...... 89

B HPLC AND ELSD DATA ...... 93

C NMR DATA ...... 94

D HRMS AND MS DATA ...... 98

E NUCLEIC ACID AND AMINO ACID SEQUENCES ...... 102

F AGAR PLATES AND C. PINENSIS CULTURES ...... 105

G TIC AND EIC DATA ...... 108

LIST OF REFERENCES ...... 111

BIOGRAPHICAL SKETCH ...... 122

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LIST OF TABLES

Table page

2-1 IC50 values for maltase by extracts of C. pinensis cultures grown in ATCC 1565 media...... 33

2-2 IC50 values for maltase by extracts of C. pinensis cultures grown in ATCC 1565 media supplemented with varying carbohydrates and polysaccharides ...... 35

2-3 IC50 values for maltase by extracts of C. pinensis cultures grown in ATCC 1565 media supplemented with glucose, chitin, glucomannan, and PLP...... 37

2-4 Reported 1H NMR chemical shifts of hydroxypyrrolidine isomers with [M + H]+= 134 ...... 38

3-1 IC50 values for maltase by extracts from heterologous expression of pCluster in E. coli ...... 55

3-2 List of deoxyoligonucleotides for expression of C. pinensis cluster and E. coli ...... 63

4-1 List of azasugar producers reported in literature ...... 68

4-2 BLASTx statistics of putative azasugar biosynthetic genes of known azasugar producers compared to azasugar biosynthetic genes in B. amyloliquefaciens ...... 70

4-3 BLASTx statistics of putative azasugar biosynthetic genes of known azasugar producers compared to azasugar biosynthetic genes in C.pin ...... 72

4-4 BLASTx statistics of putative azasugar biosynthetic genes of unknown azasugar producerscompared to azasugar biosynthetic genes in B. amyloliquefaciens ...... 74

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LIST OF FIGURES

Figure page

1-1 Diversity of glycosidase inhibitors ...... 21

1-2 Structural diversity of azasugars ...... 21

1-3 Glycosidase active site. A) Proposed oxocarbenium ion transition state. B) Proposed binding mode of deoxynojirimycin...... 22

1-4 Azasugars and analogs in market as antidiabetics or in clinical trials for treatment of lysosomal storage disorders...... 25

1-5 Proposed DNJ biosynthetic pathway in B. amyloliquefaciens FZB4222 ...... 28

1-6 Proposed biosynthetic pathway of DNJ in Commelina communis58 ...... 29

1-7 Proposed biosynthetic pathway of DMDP in Commelina communis58 ...... 29

1-8 Proposed biosynthetic pathway for nectrisine in Thelonectria discophora61 ...... 30

2-1 Homology comparison of the azasugar biosynthetic signature identified in Bacillus amyloliquefaciens FZB42 and Chitinophaga pinensis DSM 2588...... 32

2-2 Possible hydroxypyrrolidine isomers with [M + H]+ = 134 ...... 37

1 2-3 A) H NMR (300 MHz) of C. pinensis extract after SCX HPLC purification in D2O with solvent suppression.B) 1H NMR (300 MHz) of C. pinensis extract after SCX HPLC purification plus 0.5 mg standard DAB-1 in D2O with solvent suppression. C) 1 H NMR of standard DAB-1 in D2O ...... 41

2-4 A) 1H NMR (300 MHz) of C. pinensis extract after boronic acid resin purification in 1 D2O with solvent suppression. B) H NMR of standard DAB-1 in D2O ...... 41

1 1 2-5 A) H NMR (300 MHz) of standard DAB-1 in D2O. B) H NMR of C. pinensis extract after SCX HPLC in D2O with solvent suppression ...... 42

2-6 (+) ESI of standard DAB-1 (top), deuterated DAB-1 (middle), and DAB-1 isolated from C. pinensis (bottom) averaged across RT 14.6-15.8 minutes...... 42

3-1 A) pETBlue-CpinCluster plasmid used for heterologous expression in E. coli. B) SDS PAGE gel of pCluster expression in E. coli.* ...... 51

3-2 Maltase assay inhibition profile of extracts from heterologous expression of the putative azasugar cluster in E. coli after Amberlite IR120 (H+) purification ...... 52

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3-3 Comparison of time dependence maltase inhibition activity of extract from heterologous expression of putative azasugar cluster in E. coli (A) vs. extract from C. pinensis (B) post cation exchange purification ...... 53

3-4 A) Hypothesis for onset binding inhibition by inhibitor isolated from heterologous expression of putative azasugar cluster from C. pinensis in E. coli B) Reduction of iminium-moiety with NaBH4 ...... 54

3-5 Comparison of time dependence maltase inhibition activity of extract from heterologous expression of putative azasugar cluster in E. coli before and after NaBH4 reduction ...... 55

3-6 A) 1H NMR (300 MHz) of inhibitor isolated from C. pinensis B) 1H NMR of reduced inhibitor isolated from E. coli post SCX HPLC in D2O ...... 58

3-7 MS of NaBH4 reduced inhibitor isolated from heterologous expression of pCluster in E. coli post SCX HPLC ...... 58

3-8 Scheme of possible products as a result of reduction of inhibitor with NaBD4 to elucidate the chemistry of the conversion of nectrisine to DAB-1 ...... 59

3-9 HRMS of NaBD4 reduced inhibitor isolated from heterologous expression of pCluster in E. coli post SCX HPLC ...... 60

3-10 A) 1H NMR (300 MHz) of deuterated inhibitor isolated from E. coli after HILIC HPLC purification. B) 1H NMR (300 MHz) of DAB-1 standard...... 61

4-1 Dendrogram of microorganisms containing a putative azasugar biosynthetic cluster...... 67

4-2 Genome neighborhood of microorganisms with putative azasugar biosynthetic cluster and other genes of interest...... 67

5-1 Scheme of proposed biosynthetic pathway for DAB-1 in C. pinensis ...... 85

A-1 Yeast maltase assay of concentrated alkaline C. pinensis extract fractions from Amberlite IR120 (H+) purification...... 89

A-2 Almonds β-glucosidase assay of concentrated alkaline C. pinensis extract fractions from Amberlite IR120 (H+) purification...... 89

A-3 Streptomyces chitinase assay of concentrated alkaline C. pinensis extract fractions from Amberlite IR120 (H+) purification...... 90

A-4 Jack Bean mannosidase assay of concentrated alkaline C. pinensis extract fractions from Amberlite IR120 (H+) purification...... 90

A-5 Yeast maltase assay of concentrated alkaline autoclaved and non-autoclaved C. pinensis extract fractions from Amberlite IR120 (H+) purification...... 91

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A-6 Yeast maltase assay of concentrated alkaline ATCC 1565 media fractions from Amberlite IR120 (H+) purification...... 91

A-7 Jack Bean mannosidase assay of alkaline E. coli pCluster extract fractions from Amberlite IR120 (H+) purification...... 92

A-8 Yeast maltase inhibition assay of alkaline E. coli pETBlue-2 extract fractions from Amberlite IR120 (H+) purification...... 92

B-1 A) Chromatogram of C. pinensis extract after HILIC HPLC. B) Maltase inhibition profile of corresponding HILIC HPLC fractions...... 93

C-1 A) 1H NMR (500 MHz) corresponding to maltase inhibitor from C. pinensis after HILIC HPLC purification. B) 1H NMR (500 MHz) of standard D/L N-acetylarigine. .... 94

C-2 1H NMR (300 MHz) corresponding to maltase inhibitor from C. pinensis after HILIC HPLC...... 95

1 C-3 H NMR (300 MHz) corresponding to maltase inhibitor from NaBD4 reduction of inhibitor extracted from E.coi heterologous expression after SCX HPLC purification. ... 96

1 C-4 H COSY NMR (300 MHz) of NaBD4 reduced extracts, from E. coli heterologous expression, after HILIC HPLC purification ...... 97

D-1 HRMS of C. pinensis extract after HILIC HPLC Purification...... 98

D-2 HRMS of D/L- N-acetylarginine standard...... 99

D-3 HRMS of C. pinensis extract from cultures supplemented with chitin, sorbitol, and glucose after HILIC HPLC...... 100

D-4 HRMS of C. pinensis extract from cultures supplemented with chitin, glucose, and glucomannan after SCX HPLC...... 101

E-1 Cpin2154 gene sequence...... 102

E-2 Cpin2154 amino acid sequence...... 102

E-3 Cpin2153 gene sequence...... 103

E-4 Cpin2153 amino acid sequence...... 103

E-5 Cpin2152 gene sequence...... 104

E-6 Cpin2152 amino acid sequence...... 104

F-1 A) On the left, a 500 mL culture of C. pinensis in ATCC 1565 media supplemented with chitin, glucose, and glucomannan after 3 days. On the right, a 50 mL culture of

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C. pinensis in same conditions as the large culture n the left. Both have 50 µg/mL kanamycin. B) zoomed in picture of the 500 mL culture in A...... 105

F-2 ATCC 1565 agar plates of C. pinensis culture streaks from Figure F-1B...... 106

F-3 ATCC 1565 agar plates of greenish gray solid from C. pinensis cultures from Figure F-1A...... 106

F-4 ATCC 1565 agar plates of C. pinensis from frozen stock...... 107

G-1 TIC of standard DAB-1, deuterated DAB-1 isolated from heterologous expression of azasugar biosynthetic cluster in E. coli, and DAB-1 isolated from C. pinensis ...... 108

G-2 EIC of m/z 134 of standard DAB-1, deuterated DAB-1 isolated from heterologous expression of azasugar biosynthetic cluster in E. coli, and DAB-1 isolated from C. pinensis ...... 109

G-3 EIC of m/z 135 of standard DAB-1 and deuterated DAB-1 isolated from heterologous expression of azasugar biosynthetic cluster in E. coli ...... 110

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LIST OF ABBREVIATIONS

°C degrees Celsius

2AM 2-amino-2-deoxy-D-mannitol

4AD 4-amino-4-deoxyarabinitol

ACN acetonitrile

Amp ampicillin

B. amyloliquefaciens Bacillus amyloliquefaciens FZB42

Cam chloramphenicol

Carb carbenicillin

COSY homonuclear correlation spectroscopy

C. pinensis Chitinophaga pinensis DSM 2588

Da dalton

DAB-1 1,4-dideoxy-1,4-amino-arabinitol dd Doublet of doublets ddd Doublet of doublet of doublets

DGJ 1-deoxygalactonojirimycin

DMJ 1-deoxymannojirimycin

DMDP 2,5-dideoxy-2,5-imino-D-mannitol

DNJ 1-deoxynojirimycin dNTP deoxyribonucleotide triphosphate

E. coli Escherichia coli

EDTA ethylenediaminetetraacetic acid

EIC Extracted ion chromatogram

ESI Electron Spray Ionization

EtOH ethanol

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F6P fructose 6-phosphate

HILIC Hydrophilic Interaction Liquid Chromatography

HPLC High Pressure Liquid Chromatography

HRMS High Resolution Mass Spectrometry

IPTG isopropyl β-D-1-thiogalactopyranoside

Kan kanamycin

MeOH methanol

MJ mannojirimycin mg milligram min minute mL milliliter mM millimolar

MS mass spectrometry

NAD+ nicotinamide adenine dinucleotide

NADH reduced form of NAD+

ND not determined

Neo neomycin

NJ nojirimycin nmol nanomole

NMR nuclear magnetic resonance

OD600 optical density at 600 nm

PCR polymerase chain reaction

PLP pyridoxal 5′-phosphate

PNP-Glc para-nitrophenol α-D-glucopyranoside

PNP-Man para-nitrophenol α-D-mannopyranoside

14

rpm revolutions per minute s second

SCX Strong cation exchange

SDS-PAGE sodium dodecyl sulfate polyacrylamide gel electrophoresis

TIC Total ion current

TPP thiamin pyrophosphate

Tris tris(hydroxymethyl)aminomethane

U Unit

UV-vis ultraviolet to visible

WT wild type

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Abstract of Dissertation Presented to the Graduate School of the University of Florida in Partial Fulfillment of the Requirements for the Degree of Doctor of Philosophy

A FUNCTIONAL AZASUGAR BIOSYNTHETIC CLUSTER FROM CHITINOPHAGA PINENSIS

By

Claribel Nuñez

May 2019

Chair: Nicole A. Horenstein Major: Chemistry

Azasugars have been isolated from various plants and microorganisms and are reported to be potent and highly specific inhibitors of glycosidases. These enzymes are essential in many biological processes such as tumor metastasis, cell-cell regulation, and immune response. Since the discovery of azasugars in the mid-1960s, research has largely focused on their chemical syntheses and their potential implementation in both pharmaceutical and agrochemical applications.

More than 40 years after the discovery of the first azasugar deoxynojirimycin (DNJ), a signature gene cluster in the DNJ producer Bacillus amyloliquefaciens FZB42 (B. amyloliquefaciens), was identified. This cluster contains the genes gutB1, gabT1, and yktC1, which code for putative aminotransferase, phosphatase, and zinc-dependent dehydrogenase enzymes, respectively. It was determined that the GabT1 enzyme is essential for the biosynthesis of DNJ in B. amyloliquefaciens, and that the cluster is sufficient for heterologous expression of the DNJ precursor, mannojirimycin. Although azasugars display unique properties as natural products, only part of the biosynthetic pathway has been identified and the natural roles azasugars play in biological systems remain unknown. Accordingly, the work presented in this dissertation displays my efforts to uncover the unique nature of this important class of

16

compounds by identifying new azasugar producing organisms and azasugars, discovering the enzymes responsible for their biosynthesis, and ultimately providing insight into their natural roles.

In the first project, a gene cluster in Chitinophaga pinensis (C. pinensis), homologous to the B. amyloliquefaciens cluster, was identified which contained a putative aminotransferase and alcohol dehydrogenase clustered with a putative haloacid dehalogenase family member. The latter may be a phosphatase. The starting hypothesis was that C. pinensis was a DNJ producer.

Cultures of C. pinensis were grown and analyzed for α-glycosidase, β-glycosidase, chitinase, and

α-mannosidase inhibition. The inhibitor proved to be most potent towards α-glycosidases, consistent with DNJ as the active compound. The inhibitor was further purified and characterized by HRMS and NMR. An HRMS m/z peak of 134.0814 and proton NMR analysis of combined

SCX HPLC purified active fractions resulted in data consistent with the structure of theknown pyrrolidine azasugar, 1,4-dideoxy-1,4-amino-arabinitol (DAB-1), not DNJ. This provided evidence that C. pinensis produces an azasugar other than DNJ.

The second project focused on confirming that the hypothesized azasugar biosynthetic cluster in C. pinensis is responsible for the biosynthesis of DAB-1. The cluster was heterologously expressed in E. coli, a non-azasugar producing microorganism, the culture extracts were purified, and then analyzed for glycosidase inhibition. This led to the identification of nectrisine, an azasugar with potent slow tight binding α-glycosidase inhibition. Reduction of nectrisine with NaBH4 removed slow tight binding inhibition kinetics and produced an m/z peak of 134.10 when analyzed by MS, matching the mass of the previous isolated DAB-1 from C. pinensis. These results strongly support the theory that the homologous azasugar biosynthetic cluster identified in C. pinensis initiates the biosynthesis of DAB-1 by producing nectrisine,

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which can then be reduced by a putative reductase in C. pinensis to yield its final form. The results from my studies support the idea that the azasugar three gene cluster represent a general biosynthetic path leading to a number of different compounds.

Lastly, to gain some insights on the natural roles and prevalence of azasugars, bioinformatic tools were used to identify additional potential azasugar producing microorganisms. Using the Enzyme Function Initiation Enzyme Similarity Tool (EFI-EST), I identified organisms containing homologous GabT1 enzymes. Subsequently, the EFI Genome

Neighborhood Network Tool (EFI-GNT) was used to condense similarity results into organisms which contain a cluster of sugar-binding enzymes with the putative signature functions: aminotransferase, phosphatase, and dehydrogenase activities. A total of 155 unique microorganisms containing this azasugar biosynthetic signature were identified, many of which live symbiotically with plants or nematodes. It has been proposed that the production of azasugars provides a chemical defense mechanism for the symbiotes and aids in the acquiring of nutrients for survival.

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CHAPTER 1 INTRODUCTION

1.1 Introduction to Secondary Metabolites

Secondary metabolites, often referred to as natural products, are small organic molecules that have typically been isolated from plants, bacteria, algae, corals, sponges, and fungi.1 They are not essential for central metabolism, therefore their absence does not result in immediate death of the organism. Though not essential for growth, development, or reproduction, it has been hypothesized that secondary metabolites provide an advantage to these organisms by aiding in defense mechanisms, reproductive processes, and competing with other species for food.1,2

Fulfillment of these roles suggests that the activity of secondary metabolites are a consequence of their chemical structures. Evidence to support this hypothesis lies within the different kinds of organisms that make secondary metabolites, studying their production in nature, their biological activities, and analysis of the genes coding for their biosynthesis.2

The hypothesis that secondary metabolites are produced as part of a strategy for survival—serving as an alternative defense mechanism—stems from the observation that they are most commonly produced in organisms that lack an immune system. In plants, they serve as pesticides and protect against pathogens through their antibiotic and antifungal properties.2 Many compounds have also been shown to have anti-viral and anti-tumor effects. Somebehave as immunosuppressants and others, for example, have the ability to lower cholesterol.1,2

Unlike primary metabolites, secondary metabolites belong to more genetically complex pathways that are energetically expensive. A significant portion of an organism’s genome is often dedicated to encoding necessary information for the biosynthesis of secondary metabolites, which strongly suggests that they have been evolutionarily selected by the organism and that the metabolites are not an artifact of isolation procedures. This is further evidenced by the clustering

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of microbial genes, suggesting that the genes evolved as a unit, facilitating the simultaneous expression of the enzymes involved in the biosynthetic pathway.2,3

Classical methods for the identification and isolation of significant secondary metabolites include monitoring by bioactivity assays, or antioxidant and cytotoxicity activities that yield active principles.4 With the growing availability of genome sequences, identification of gene clusters that are homologous to others of known function has also been used in conjunction with bioassay guided techniques for the identification and characterization of secondary metabolites and the identification of new natural sources of production. In this dissertation, I describe the work aimed at identifying new sources that produce secondary metabolites known as azasugars and the enzymes responsible for their biosynthesis in an effort to begin to understand their role in nature.

1.2 Glycosidase Inhibitors

Glycosidases are ubiquitous enzymes that break down oligosaccharides and glycoconjugates by catalyzing the hydrolysis of glycosidic linkages. In addition to their roles in healthy human metabolism that include dietary assimilation of carbohydrates and in the trimming reactions of glycoconjugates that occur in the endoplasmic reticulum and Golgi apparatus,5,6 they play a key role in many metabolic disorders and human diseases such as type II diabetes, lysosomal storage disease, cancer, and viral infections.7–14 Treatment for some of these illnesses sometimes involve use of naturally occurring glycosidase inhibitors. Many different kinds of natural glycosidase inhibitors have been extracted from plants15–18 and microorganisms.19–22 The various types of glycosidase inhibitors isolated include azasugars (1), thiosugars (2), carbaglycosylamines (3), and polyhydroxylated alkaloids (4), Figure 1-1.17,23–26 For this dissertation, the focus will lie on the azasugar glycosidase inhibitors.

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Figure 1-1. Diversity of glycosidase inhibitors

1.3 Azasugars

1.3.1 Overview

Azasugars is a term often used loosely to include five classes based on structural features: piperidines, pyrrolidines (4), pyrrolizidines (8), indolizidines (6), and nortropanes (7) (Figure 1-

2). Often, there is overlap in the classification, especially with the alkaloids. Piperidines and pyrrolidines are monocyclic alkaloids, while pyrrolizidines, indolizidines, and nortropanes are

Pyrrolidines Piperidines Indolizidines

(5) (6) (4)

Pyrrolizidines Nortropanes

(7) (8)

Figure 1-2. Structural diversity of azasugars

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bicyclic heterocycles. Hydroxylated piperidines are also considered pyranose mimics, and pyrrolidines resemble furanoses because of their five-membered ring.

Azasugars are often referred to as “sugar mimics” due to their structural resemblance to sugar molecules. Azasugars, capable of bearing a positive charge at the nitrogen, mimic the oxocarbenium-like character proposed in the transition states of enzymatic glycoside hydrolysis mechanisms (Figure 1-3A).27 The nitrogen in the ring is considered to be the most relevant characteristic of azasugars for enzyme inhibition, because they feature a secondary amine that when protonated ,resembles the positive charge present in the oxocarbenium ion like transition states of glycosyl hydrolysis reactions, (Figure 1-3B).28 The hypothesis that the protonated amine is a major contributor for the inhibition of glycosidases is strongly supported by crystallographic studies on isofagomine azasugars derivatives.29

While there are many applications for glycosidase inhibitors (discussed in section 1.3.2), their natural function is not well characterized. There is no current data on the exact mechanism of action of the glycosidase inhibitors produced by microorganisms; however, there are a few hypotheses based on the observed functions of these compounds. One of the leading hypotheses for its production in plants postulates that these naturally occurring glycosidase inhibitors

(A) (B)

Figure 1-3. Glycosidase active site. A) Proposed oxocarbenium ion transition state. B) Proposed binding mode of deoxynojirimycin.

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function as antifeedants.26,30,31 Other hypotheses include the idea that microorganisms such as strains of Bacillus, Streptomyces and Metarhizium, may produce these compounds to hinder competing microorganisms from obtaining their food source by inhibiting the competing species’ glycosidases.32,33

1.3.2 Therapeutic Potential and Applications of Azasugars

1.3.2.1 Azasugars as antidiabetic agents

Due to their potent inhibition of glycosidases, azasugars have been explored as therapeutic targets for these enzymes. The brush-border membrane of the small intestine contains

α-glucosidase enzymes that digest carbohydrates into monosaccharides for improved intestinal absorption. Glucose is the main product formed as a result of these α-glucosidase-mediated hydrolysis. By inhibiting these enzymes, azasugars can help reduce the amount of sugar released into the blood, making them useful antidiabetic drugs. In 1996, an analog of DNJ (5), Miglitol

(Figure 1-4), was approved by the FDA and introduced into the market in 1999 for the treatment of Type 2 Diabetes.34 A consequence of this therapy for diabetes is that some inhibition of non- target intestinal glycosidases results in gastrointestinal discomfort because of undigested carbohydrates.23,35

1.3.2.2 Azasugars in lysosomal storage diseases

Experimental data has shown that some human genetic diseases are due to mutations in proteins that influence their folding and lead to the retention of these mutated proteins in the endoplasmic reticulum (ER), leading to degradation.36 Such genetic diseases include some lysosomal storage diseases such as Fabry, Gaucher, and Pompe diseases. In Fabry patients, studies on mutant α-galactosidase A (α-Gal A) activity revealed that the enzymes had similar

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kinetics as wild type α-Gal A, but were significantly less stable.37 In addition, the α-Gal A mutants formed aggregates in the ER leading to quick degradation of the mutants.38

Studies on the effects of azasugars on the activity of α-Gal A revealed that 1- deoxygalactonojirimycin (DGJ, 10) (Figure 1-4) enhanced the activity of mutant enzymes from

Fabry patients when administered at sub-inhibition quantities (concentration levels of the azasugar that do not produces enzyme inhibition).8 Similarly, in Gaucher disease, a lysosomal storage disorder caused by lack of β-glucocerebrosidase activity, when the azasugar, N-nonyl-

DNJ (NN-DNJ, 12) (Figure 1-4) was administered at sub-inhibitory concentrations activity of the mutant enzyme increased by two-fold.39 Studies have also shown that treatment with DNJ N- butyl-1-deoxygalactonojirimycin (NB-DGJ, 11) (Figure 1-4) lead to up to a 7.5-fold increase or

5.6-fold increase of residual mutant acid α-glucosidase (GAA), respectively, in patients with

Pompe disease.40 With respect to these lysosomal storage diseases, the azasugars are not considered to act as glycosidase inhibitors; instead they are thought to act as pharmacological chaperones. The inhibitors bind to an allosteric site of the mutant enzyme, acting as template that stabilizes the native folding state in the ER. By stabilizing the mutant enzyme and allowing it to fold correctly, the mutant can then be successfully trafficked to the lysosome and restore the intracellular activity of the mutant enzymes.39 Various azasugars and their analogs are currently going through clinical trials as pharmacological chaperones for the treatment of lysosomal storage diseases.39,41–43

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(9) (10) (11)

(5) (12)

Figure 1-4. Azasugars and analogs in market as antidiabetics or in clinical trials for treatment of lysosomal storage disorders.

1.3.2.3 Additional potential therapeutic applications of azasugars

In addition to the medicinal properties mentioned above, azasugars also possess the potential to serve as anti-viral and anti-cancer drugs. Several studies have shown compelling evidence that azasugars inhibit the spread of the human immunodeficiency virus (HIV).44,45 The virus has an envelope glycoprotein, gp120, which is heavily N-glycosylated. This glycoprotein is one of the major components that gives HIV its ability to bind to helper T-cells and induce its viral infectivity.12 By interfering with the glycosylation process of these viral glycoproteins, azasugars can reduce the activity of the virus. Similarly, the envelope glycoproteins of hepatitis

B virus (HBV) have two glycosylation sites that are sensitive to inhibitors of the N-linked glycosylation pathway.46 When treated with NB-DNJ (11), in vitro, it was found that a high proportion of the HBV virus was retained inside the cells.47 These results suggest that the glycosylation processes are necessary for the successful transport of the virus out of the host cells.

25

In cancer, aberrant glycosylation patterns are directly related to tumor cell invasion and metastasis.48 Azasugars could serve as cancer chemotherapeutics by inhibiting the catabolic and processing glycosidases involved in oncogenic transformations. A myriad of naturally occurring azasugars have been shown to exhibit anti-cancer and anti-metastatic activity by inhibiting the aggregation and adhesion of tumor cells.49 Regardless of an azasugar’s interesting in vitro properties as potential anti-viral and chemotherapeutics, the adverse effects that they produce as

α-glucosidase inhibitors has significantly hindered development of their applications.50,51 Current research has been focused on increasing bioavailability and avoiding off-target interactions with gastrointestinal glucosidases by designing and synthesizing azasugar prodrugs.52–54

1.3.3 Biosynthetic Pathways of Azasugars

Since their isolation in the mid 1960’s, research has largely focused on the chemical syntheses of azasugars and their potential implementation in both pharmaceutical and agrochemical applications. It was not until the early 1990’s that isotope labeling experiments revealed that glucose is a precursor to DNJ (5) and in 2011 a three gene cluster involved in the initiation of DNJ biosynthesis was identified.22,55

1.3.3.1 Biosynthesis of nojirimycins

The azasugars nojirimycin (NJ, 18, Figure 1-5) and DNJ (5), have been isolated from a variety of Streptomyces and Bacillus strains, as well as plants including Morus alba and

Commelina communis.56–58 Feeding experiments with glucose deuterated at C-1 or C-2 led to the isolation of deuterated DNJ at the C-6 position, suggesting that the first step in their biosynthesis is the isomerization of glucose to fructose.55 Using C-1 13C-labeled glucose led to the production of C-6 labelled DNJ, indicating that a head to tail inversion of the carbon chain must have

26

occurred.20 Deuterium isotope labeling experiments also revealed that oxidation occurs at the 6- position of the glucose/fructose compound prior to the head to tail carbon chain inversion. This lead to the biosynthetic pathway proposal in which glucose is isomerized to fructose, the C2 position is then aminated, followed by oxidation of the C6 hydroxyl group. The oxidation of the hydroxyl group causes the spontaneous cyclization that leads to the formation of the azasugar mannojirimycin (MJ, 17, Figure 1-5). Epimerization at the C2 position of MJ would produce NJ, followed by a dehydration and reduction at the C1 position to produce the final azasugar, DNJ.

Using the proposed biosynthetic pathway for DNJ as a guide, the Horenstein group then used the reported22 genome sequence for Bacillus amyloliquefaciens (B. amyloliquefaciens)

FZB42, a known DNJ producer, to identify the genes that may be responsible for its biosynthesis. A cluster of three (putative) sugar binding genes coding for a transaminase

(gabT1), a phosphatase (yktC1), and an alcohol dehydrogenase (gutB1) were identified.

Considering the findings of Hardick et al and the newly identified cluster, it was proposed that glucose entered glycolysis to become fructose-6-phosphate (F6P, 13, Figure 1-5), which is aminated by GabT1 to form 2-amino-2-deoxy-D-mannitol-6-phosphate (2AM6P, 14, Figure 1-

5). Then 2AM6P is dephosphorylated by YktC1 into 2-amino-2-deoxy-D-mannitol (2AM, 15,

Figure 1-5) which is oxidized by GutB1 to form an intermediate 6-oxo-compound (16) that spontaneously cyclizes to MJ. Finally, MJ is converted to DNJ by an epimerization and reduction step (Figure 1-5).22

Knockout of the gabT1 gene in B. amyloliquefaciens completely abolished DNJ production. Growth in the presence of 2AM6P to the B. amyloliquefaciens gabT1 mutant reinstated the production of DNJ. In addition, expression of the gene cluster in E. coli, a bacterium that does not produce azasugars, led to the isolation of MJ.22,59 In another case, it was

27

(13) (14)

(15) (16)

(17) (18) (5)

Figure 1-5. Proposed DNJ biosynthetic pathway in B. amyloliquefaciens FZB4222

found that the three gene cluster from Bacillus subtilis MORI 3k-85 was responsible for DNJ production when expressed in E. coli,60 implying that the three genes were sufficient for its biosynthesis, or E.coli provided the rest of the required enzymes. Expression of the three-gene cluster from B. subtilis MORI 3k-85 in E. coli resulted in the isolation of DNJ, which was confirmed by LC-MS/MS. These experiments provided evidence that the three gene cluster was indeed responsible for the initiation of the biosynthesis of DNJ. The remaining enzymes involved in the DNJ biosynthetic pathway remain to be elucidated.

A different biosynthetic route has been observed for DNJ in plants such as Commelina communis. Carbon labeled glucose at the C-1 position led to DNJ with 13C enrichment located at the C-1 instead of the C-6 position.58 This led to a proposal in which a C-1/C-5 cyclization is produced in the original glucose molecule without any type of inversion (Figure 1-6). The same

13C enrichment at C-1 for fructose was observed when C-1 carbon labeled glucose was administered to the plant.58

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Figure 1-6. Proposed biosynthetic pathway of DNJ in Commelina communis58

1.3.3.2 Biosynthesis of DMPD

From the same labeling experiments in Commelina communis that identified a C-1/C-5 cyclization for DNJ production, the same labeling patterns were observed for DMDP, another prominent azasugar made by this plant.58 The proposed biosynthetic pathway is depicted in

Figure 1-7.

Figure 1-7. Proposed biosynthetic pathway of DMDP in Commelina communis58

1.3.3.3 Biosynthesis of nectrisine

Most recently, the biosynthetic genes for the pyrrolidine azasugar nectrisine was identified in the fungus Thelonectria discophora SANK 18292.61–63 Feeding the fungus carbon

29

labeled D-ribose and D-xylose, produced carbon labeled nectrisine, suggesting that both these compounds are precursors. This led to the proposal that the fungus utilizes the pentose phosphate pathway for the biosynthesis of nectrisine. Like the DNJ producing microorganisms,

Thelonectria discophora was identified as having a three gene cluster coding for proteins that had sugar binding motifs, consisting of an aminotransferase (necA), phosphatase (necB), and an oxidase (necC).61 Expression of this three gene cluster in E. coli was sufficient for the biosynthesis of nectrisine.61 Based on these experiments the following biosynthetic pathway for nectrisine was proposed: D-ribose or D-xylose gets converted to xylulose-5-phosphate through the pentose phosphate pathway, which then gets converted to 4-amino-4-deoxyarabinitol. It is unclear whether dephosphorylation or amination happens first. NecC then oxidizes 4-amino-4- deoxyarabinitol to form the azasugar nectrisine (Figure 1-8).

Figure 1-8. Proposed biosynthetic pathway for nectrisine in Thelonectria discophora61

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CHAPTER 2 IDENTIFICATION AND CHARACTERIZATION OF DAB-1 IN C. PINENSIS

2.1 Overview

The gene sequences for the azasugar biosynthetic signature identified in B. amyloliquefaciens was used to identify organisms with a homologous cluster and thus, putative azasugar production using pBLAST (protein Basic Local Alignment Sequence Tool) from the

National Center for Biotechnology Information (NCBI) and EFI-GNT (Enzyme Function

Initiative- Genome Neighborhood Network Tool). One such organism was the terrestrial gliding bacterium Chitinophaga pinensis (C. pinensis).64 Given its interesting chitinolytic properties and low laboratory biosafety level, we chose to work with C. pinensis to identify and characterize its potential for azasugar production. The identified azasugar biosynthetic cluster in C. pinensis consists of homologous putative aminotransferase and alcohol dehydrogenase clustered with a putative haloacid dehalogenase family, which include phosphatases (Figure 2-1). We originally hypothesized that C. pinensis would produce DNJ.

In this chapter, evidence is provided showing that C. pinensis does in fact produce a pyrrolidine azasugar, 1,4-dideoxy-1,4-imino-arabinitol (DAB-1). To our knowledge, there is no bacterial source reported in the literature for the biosynthesis of DAB-1, a potent α-glycosidase inhibitor previously isolated from plants.65,66 Maximum inhibitor production was observed after culturing bacteria at room temperature for three days in liquid media supplemented with glucose, chitin, and glucomannan. After various purification steps the structure of DAB-1 was elucidated by HRMS and proton NMR.

31

Haloacid Dehalogenase

Figure 2-1. Homology comparison of the azasugar biosynthetic signature identified in Bacillus amyloliquefaciens FZB42 and Chitinophaga pinensis DSM 2588.

2.2 Results and Discussion

First, we wanted to determine if C. pinensis produced a glycosidase inhibitor, specifically an azasugar. Initially, cultures of C. pinensis were grown in the suggested ATCC 1565 growth media and the extract was partially purified by cation exchange chromatography on an Amberlite

IR 120 (H+) column eluted with water and then aqueous ammonium hydroxide. The alkaline fractions of different C. pinensis cultures were analyzed for α-glucosidase (Figure A-1), β- glucosidase (Figure A-2), chitinase (Figure A-3), and α-mannosidase (Figure A-4) inhibition.

The inhibitor was most potent towards α-glucosidase; therefore, the maltase assay was used to search for fractions containing the inhibitor in the remaining purification steps. It is important to note that in some experiments, the C. pinensis cultures were harvested and autoclaved to kill the bacteria. Control experiments were run to ensure that the inhibition that was observed was not an artifact caused by autoclaving (Figure A-5). Even though autoclaving did not affect inhibition, later experiments were not autoclaved to avoid possible impurities caused by degradation of complex structures as a result of autoclaving the C. pinensis cultures. To further confirm that the

32

inhibition was being produced by a C. pinensis inhibitor and not an artifact of the growth media, the ATCC 1565 media was loaded onto a cation exchange column and the alkaline fractions were analyzed for maltase inhibition.

After confirmation that the inhibition was not due to an artifact, fractions possessing maltase inhibition from the cation exchange column were then loaded onto an anion exchange column followed by HILIC HPLC. The apparent IC50 value of the C. pinensis extracts with maltase after each purification step was calculated (Table 2-1). The HILIC HPLC chromatogram,

1H NMR, and HRMS of the partially purified inhibitor after HILIC HPLC are in the Appendix

Figures B-1, C-1, and D-1, respectively. From these preliminary data we observe a few things: after HILIC HPLC an IC50 value of 700 µg/mL is obtained, which is indicative of the presence of a glycosidase inhibitor. Also, the IC50 values decrease after each purification step; a 17-fold increase in inhibition after HILIC HPLC compared to inhibition activity after the cation exchange column is noted. This increase in inhibition potency suggests that the inhibitor was being successfully purified since less of the extract was required to inhibit 50 % of the enzyme.

Considering that we were trying to identify whether C. pinensis produces an azasugar, and azasugars have been reported to have glycosidase inhibition potency in the nanomolar65,66 and

57,67 low micromolar scales, this inhibitor with a 700 µg/mL IC50 value that was isolated did not seem promising.

Table 2-1. IC50 values for maltase by extracts of C. pinensis cultures grown in ATCC 1565 media Purification Method Total (g) IC50 (μg/mL) Cation Exchange 1.83 12000 Anion Exchange 0.83 7000 HILIC HPLC 0.013* 700 The total is the mass of the combined C. pinensis extract fractions that showed maltase inhibition after each of the purification method. The IC50 value is the amount of C. pinensis extract added to reaction mixture that inhibited maltase by 50 %. *The total mass for HILIC HPLC is a combination of inhibitory fractions of C. pinensis extract from multiple HPLC runs.

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However, this apparent weak inhibition was most likely due to impurities that remained in the sample, as can be inferred from the impure HPLC chromatogram (Figure B-1A) and

HRMS data (Figure D-1). In addition, after HILIC HPLC purification, the major product that can be seen in the NMR appears to be N-acetylarginine (Figure C-1), which is not an inhibitor of yeast maltase. The HRMS of N-acetylarginine was also obtained to compare with the HRMS of the partially purified C. pinensis extracts (Figure D-2). These results imply that the quantity of the inhibitor being produced and isolated was minimal and therefore not enough for characterization. Consequently, our efforts then shifted to optimization of growth conditions for maximum inhibitor production. C. pinensis growth cultures were supplemented with either 1 % glucose and 1 % sorbitol or 0.1 % chitin. After harvesting, the culture extracts were purified and evaluated for maltase inhibition as described previously.

Supplementing the C. pinensis cultures with glucose and sorbitol led to IC50 values of less than 1 μg/mL after HILIC HPLC purification (Table 2-2), a 700-fold increase compared to culture grown in ATCC 1565 media alone. Further, supplementing with glucose, sorbitol, and chitin led to an IC50 value of 5µg/mL after the anion exchange column purification (Table 2-2), an 800-fold increase in inhibition compared to extracts from cultures grown in ATCC 1565 media, which consists of casitone as its carbon source, after anion exchange column purification.

The maximum inhibition calculated for the extracts were now within the range of the IC50 values of 0.33 - 2.6 μg/mL reported in the literature for DNJ. 57,67 The sample was submitted for HRMS, but there was not enough to obtain an NMR. Given that the major peak corresponded to N- acetylarginine (Figure D-3), it was determined that the purification methods being used was not adequate for the removal of the persistent N-acetylarginine impurity; additional steps needed to be implemented.

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Table 2-2. IC50 values for maltase by extracts of C. pinensis cultures grown in ATCC 1565 media supplemented with varying carbohydrates and polysaccharides ATCC 1565 Media ATCC 1565 Media supplemented with 1 % supplemented with 1 % Purification Method Glucose and 1 % Sorbitol Glucose, 1 % Sorbitol 0.1 %, per 1L and Chitin per 1L IC IC Total (g) 50 Total (g) 50 (μg/mL) (μg/mL) Cation Exchange Column 1.58 160 2.98 100 Anion Exchange Column 0.170 50 0.1367 5 HILIC HPLC ≤0.001 ≤1 ND ND

This table shows the maltase IC50 values of C. pinensis extracts of cultures that were supplemented with 1 % glucose and 1 % sorbitol or 0.1 % chitin. The total is the mass of the combined C. pinensis extract fractions that showed maltase inhibition after each of the purification method. The IC50 value is the amount of C. pinensis extract added to reaction mixture that inhibited maltase by 50 %. The minimum mass that the balance can measure is 1 mg, and so this was used as the mass value for extract mass that was below detection limit to estimate the IC50. ND= not determined

Different phenotypes were observed in C. pinensis cultures, which led to concerns that the cell cultures may have been contaminated. Not much information can be found about C. pinensis in literature except for the antibiotic resistance profile and basic growth details and conditions.64,68,69 The literature only reports the growth of C. pinensis as producing a yellowish pigment. By contrast, when we worked with C. pinensis cultures, we observed yellow, orange, and greenish yellow pigments. On the greenish yellow cultures, a grey residue was observed on the walls of the flasks (Figure F-1). To ensure that the yellow greenish cultures were C. pinensis with different phenotypes and not a result of contaminants, the cultures and the residue were re- streaked on ATCC 1565 agar plates with no antibiotics, 50 μg/mL kanamycin (kan), 50 μg/mL carbenicillin (carb), or 10 μg/mL neomycin (neo), (Figures F-2 and F-3). Control plates of C. pinensis from frozen stocks were also prepared (Figure F-4). In addition, although C. pinensis is a gliding bacterium, colonies can be obtained when using a serial streak dilution technique; streaking cells on an agar plate and then picking cells from the streaked area and re-streaking in

35

another section of the plate (Figures F-2, F-3, and F-4). The single colonies sometimes did not grow when used to start liquid cultures and their life span on plates when stored at 4 ºC was shorter than that of C. pinensis colonies that were clustered into a film.

In further inhibitor isolation experiments, C. pinensis cultures were grown in ATCC 1565 media supplemented with chitin, glucose, and glucomannan. The glucomannan was added because it has been shown to support the greatest growth for C. pinensis.68 In order to reduce the amount of impurities, C. pinensis cell pellets were resuspended in minimal media supplemented with chitin, glucose, and glucomannan. In addition, 50 μg of PLP was sterile filtered and added to the cultures increase the expression of the putative PLP-dependent aminotransferase. The cultures were harvested and purified by the following methods: cation exchange column eluted with 0.5 M NH4OH, followed by methanol extraction, anion exchange column eluted with dI

H2O, C18 HPLC with isocratic elution of 1 % acetonitrile (ACN), and finally strong cation exchange (SCX) HPLC with a gradient elution of 1 % 50 mM ammonium acetate (NH4Ac) followed by 30 % 50 mM NH4Ac. The extracts were analyzed for maltase inhibition after each purification method and then analyzed by 1H NMR and HRMS.

Growth of C. pinensis cultures in minimal media supplemented with chitin, glucose, glucomannan, and PLP led to the isolation of a potent α-glycosidase inhibitor (Table 2-3). The presence of a [M + H]+ = 134.0814 peak in the HRMS chromatogram (Figure D-4) suggested the presence of a pyrrolidinic azasugar (Figure 2-2). Iminopentitols possessing [M + H]+ = 134 mass have been previously isolated from plants and marine sponges.70,71 Hydroxypyrrolidine isomers have also been synthesized (Table 2-4) and surveyed for glycosidase inhibition.72–76 In the proton

NMR of the inhibitor isolated from C. pinensis after SCX HPLC, a doublet of doublets (dd) at

3.24 ppm with J coupling constants of 12.6 and 2.4 Hz was observed (Figure 2-3A). In

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Table 2-3. IC50 values for maltase by extracts of C. pinensis cultures grown in ATCC 1565 media supplemented with glucose, chitin, glucomannan, and PLP. IC Dry Mass Weight from 30 L of Culture Purification 50 (μg/mL) (mg) Cation Exchange Column 1160 580 MeOH Extraction 780 390 Anion Exchange Column 6 30 C18 HPLC ≤2 <1 SCX HPLC ≤2 <1 HILIC HPLC ND ND This table shows the maltase IC50 values of C. pinensis extracts of cultures supplemented with glucose, chitin, PLP, and glucomannan. The total is the mass of the combined C. pinensis extract fractions that showed maltase inhibition after each of the purification method. The IC50 value is the amount of C. pinensis extract added to reaction mixture that inhibited maltase by 50 %. The minimum mass that the balance can readily measure is 1.0 mg, and so this was used as the maximal mass value for extract mass that was below detection limit to estimate the IC50. ND= not determined

addition, an overlapping dd that resembles a triplet was observed at around 3.98 ppm with J coupling constants of 3.8 and 3.2 Hz. A multiplet with J couplings of 4.7, 3.1 and 2.4 Hz was also observed at 4.22 ppm. If we compare these proton peaks and their J coupling constants to the reported proton NMR and J coupling constants of the ribo, lyxo, xylo, and arabino isomers reported in literature (Table 2-4), it can be observed that the inhibitor is most like the arabino isomer; it contains a multiplet, which they label as ddd, with J coupling constants of 4.5, 3.3, and

2.4 Hz at 4.26 ppm, a dd at 3.29 with J coupling constants of 12.6 and 2.4 Hz, and a dd at 4.02 ppm with J coupling constants of 3.6 and 3.3 Hz.75,77 The proton NMR data available shows that

Figure 2-2. Possible hydroxypyrrolidine isomers with [M + H]+ = 134

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Table 2-4. Reported 1H NMR chemical shifts of hydroxypyrrolidine isomers with [M + H]+= 134 δ, ppm, mult. (J, Hz)ref H- position Ribo75,76 Lyxo75 Xylo73,75 Arabino75,77 1a 3.43, dd (12.9, 3.9) 3.48, dd (12.0, 7.8) 3.60, dd (12.9, 4.2) 3.51, dd (12.6, 4.5) 1b 3.30, dd (12.9, 1.8) 3.15, dd (12.0, 3.9) 3.23, broad d (12.9) 3.29, dd (12.6, 2.4) 2 4.31, ddd (3.9, 3.9, 1.8) 4.44, ddd (7.8, 4.8, 3.9) 4.32, broad d (3.3) 4.26, ddd (4.5, 3.3, 2.4) 3 4.13, dd (8.7, 3.9) 4.29, dd (4.2, 3.9) 4.25, broad s 4.02, dd (3.6, 3.3) 4 3.56, ddd (8.7, 6.0, 3.3) 3.69, ddd (8.2, 4.8, 4.2) 3.55, ddd (8.1, 5.7, 3.9) 3.83, m 5a 3.75, dd (12.3, 6.0) 3.84, dd (12.0, 8.2) 3.76, dd (12.3, 8.1) 5b 3.90, dd (12.3, 3.3) 3.93, dd (12.0, 4.8) 3.95, dd (12. 4.2) 3.88, dd, (12.0, 5.7)

the inhibitor produced by C. pinensis is most likely the azasugar 1,4-dideoxy-1,4-amino- arabinitol (DAB-1, 19) or it’s L enantiomer (LAB-1): the proton in both L and D isomers have the same chemical shift.75,77,78

DAB-1 and LAB-1 are both potent inhibitors of the yeast maltase enzyme. The L enantiomer, however, has a slightly lower IC50 value towards maltase and, unlike DAB-1, does not inhibit β-glucosidase.72 The inhibitor isolated from C. pinensis inhibits β-glucosidase to a lesser extent compared to maltase (Figure A-2) and therefore it’s identity is most likely DAB-1.

Comparison of the 1H NMR of the inhibitor from C. pinensis after SCX-HPLC purification and standard DAB-1 is shown in Figure 2-3A. We hypothesized that the impurity observed in the 1H

NMR of the inhibitor isolated from C. pinensis may be the aminopolyol 4-amino-4- deoxyarabinitol (4AD, 20, Figure 2-3A). The presence of 4AD was supported by the second largest peak of 152.0916 in the HRMS (Figure D-4). For further confirmation of the azasugar’s stereochemistry, the azasugar isolated from C. pinensis and the DAB-1 standard can be derivatized using Marfey’s reagent and analyzed by HPLC.71 Comparison of the retention time can then confirm if the azasugar from C. pinensis is DAB-1.

To support the hypothesis that DAB-1 was present in the sample by NMR, 0.5 mg of standard DAB-1 was added to the sample in Figure 2-3A. The idea was that by adding a small amount of standard DAB-1 there would be an increase in peaks related to DAB-1, if present,

38

providing evidence for the presence of DAB-1 in the extract. If the addition of standard DAB-1 gave new peaks with slightly different chemical shifts, then it would provide evidence for a different molecule with some structural similarities to the DAB-1. Unfortunately, the addition of

0.5 mg, although a small amount in routine NMR, was too much for the supposed DAB-1 in the sample; it was difficult to determine if there was any difference to original sample because the impurities that were clearly visible were now part of the baseline (Figure 2-3B). The now compromised sample was then used to develop methods for the separation of DAB-1 from the remaining impurities. To separate DAB-1 from 4AD, the compromised mixture was loaded onto a boronic acid resin column, eluted with dI H2O, and fractions were assayed for maltase inhibition. 1H NMR of combined fractions exhibiting maltase inhibition is shown in Figure 2-4A.

The NMR shows that 4AD was successfully separated from DAB-1. The difference in chemical shifts can be due to lower concentration of DAB-1, as a result of sample loss during purification, and change of lock signal position caused by different ionic strength of the samples.

Since standard DAB-1 was added to the original sample, more sample was prepared by growing C. pinensis cultures as described before. The culture supernatant was purified and analyzed for maltase inhibition as previously described. A HILIC HPLC purification step was added prior to SCX HPLC to the protocol in this round due to additional impurities that were found in this sample (Figure C-2). After SCX HPLC, we observed evidence for the presence of

DAB-1 in the 1H NMR (Figure 2-5). However, a very small amount of inhibitor was isolated, and we were unable to remove all the impurities which made the sample difficult to analyze by

NMR. To provide evidence that this sample does in fact contain DAB-1, the sample along with standard DAB-1 and the deuterated DAB-1 obtained in Chapter 3 was submitted to the

University of Florida Department of Chemistry Mass Spectrometry Research and Education

39

Center for LC-MS analysis. The total ion current (TIC) chromatogram of the samples are in the

Appendix figure G-1. The extracted ion current (EIC) chromatograms for m/z 134 and m/z 135 can be found in the Appendix, Figures G-2 and G-3 respectively. The positive electron spray ionization MS averaged across RT 14.6 – 15.8 minutes can be found in Figure 2-6. The LC-MS data shows that all three samples, standard DAB-1, deuterated DAB-1 isolated from heterologous expression of the putative azasugar biosynthetic cluster in E. coli, and inhibitor isolated from C. pinensis extracts eluted at the same time showing their corresponding 134 and

135 [M + H]+ peaks. This provides evidence that the inhibitor being isolated from C. pinensis is

DAB-1 and DAB-1 is the molecule that is obtained after reduction of the inhibitor isolated from heterologously expression the cluster in E.coli (further discussed in Chapter 3).

In summary, I provide evidence that C. pinensis produces an azasugar, but not DNJ. The maltase assay guiding the isolation would have been appropriate to detect DNJ if it were present in significant quantity. The structure of the azasugar isolated from C. pinensis matches that of

DAB-1, a known azasugar produced in plants and fungus. This is the first known source for bacterial production of DAB-1, and the work provides the first evidence that the three gene signature azasugar cluster in bacteria can code for biosynthesis of compounds other than DNJ.

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C)

(19) DAB-1 B)

A)

(20) (19) 4AD DAB-1

1 Figure 2-3. A) H NMR (300 MHz) of C. pinensis extract after SCX HPLC purification in D2O with solvent suppression.B) 1H NMR (300 MHz) of C. pinensis extract after SCX HPLC purification plus 0.5 mg standard DAB-1 in D2O with solvent suppression. C) 1 H NMR of standard DAB-1 in D2O

A)

B) (19) DAB-1

Figure 2-4. A) 1H NMR (300 MHz) of C. pinensis extract after boronic acid resin purification in 1 D2O with solvent suppression. B) H NMR of standard DAB-1 in D2O

41

A)

B)

1 1 Figure 2-5. A) H NMR (300 MHz) of standard DAB-1 in D2O. B) H NMR of C. pinensis extract after SCX HPLC in D2O with solvent suppression

Standard DAB-1

Deuterated DAB-1

DAB-1 from C. pinensis

Figure 2-6. (+) ESI of standard DAB-1 (top), deuterated DAB-1 (middle), and DAB-1 isolated from C. pinensis (bottom) averaged across RT 14.6-15.8 minutes.

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2.3 Experimental

2.3.1 General Methods

Chitinophaga pinensis strain UQM 2034 was obtained from ATCC. Growth media components bacto-casitone, magnesium sulfate, yeast extract, sodium phosphate, potassium phosphate, ammonium chloride, sodium chloride, calcium chloride, glucose, and sodium hydroxide, as well as ammonium hydroxide, acetic acid, 195 proof ethanol and hydrochloric acid were purchased from Fisher Scientific. Other growth media components such as Agar, kanamycin, neomycin, and chitin were purchased from Sigma Aldrich. Carbenicillin was purchased from Boston BioProducts. The Amberlite IR 120 (H+) and Dowex 1x8 200-400 mesh

(Cl-) used for purification were purchased from Alfa Aesar. The polymer bound boronic acid resin, 200-400 mesh, was purchased from Sigma Aldrich. For inhibition assays, yeast maltase was purchased from Fluka analytical, β-glucosidase from almonds, chitinase from Streptomyces,

Jack Bean mannosidase, and their respective p-nitrophenyl glycopyranosides substrates, and chitin azure were purchased from Sigma Aldrich. Purified samples isolated from C. pinensis were sent for MS and/or HRMS analysis at UF’s Mass Spectrometry Research and Education

Center.

2.3.2 Growth Conditions for Chitinophaga pinensis and Culture Extract Purification

2.3.2.1 Rich media growth conditions

Chitinophaga pinensis was propagated on ATCC 1565 (5 g casitone, 3 g yeast extract, 1 g MgSO4•7H2O per 1 L) media plates supplemented with 50 g/mL kanamycin. Starter cultures

(5 mL) grown in ATCC 1565 overnight at 25 °C were inoculated into 1 L ATCC 1565 media supplemented with 3 g of glucomannan, 1 g chitin, 10 mL of a 50 % sterile filtered glucose solution, 50 g sterile filtered PLP and 50 g mL-1 kanamycin. The cultures were grown at 25

43

C, with shaking at 250 rpm for 3 days. The cultures were then centrifuged in a GS 3 rotor at

6000 rpm at 4 C for 40 minutes to remove the cell pellet. The supernatant was collected, purified and screened for glycosidase inhibition.

2.3.2.2 Minimal media growth conditions

A Chitinophaga pinensis starter culture was propagated in rich media conditions as described above. After overnight inoculation in ATCC 1565 at 25 °C, the 1 L cultures were centrifuged in a GS 3 rotor at 6000 rpm at 4 C for 40 minutes. The supernatant was discarded, and the cells were resuspended in 250 mL M9 minimal media (64 g Na2HPO4.7H2O, 15 g

KH2PO4, 5 g NH4Cl, 2.5 g NaCl, 2 mM MgSO4, 0.1mM CaCl2 per L at pH 7) supplemented with

1.5 g of glucomannan, 0.5 g chitin, 5 mL of a 50 % sterile filtered glucose solution, 50 g sterile filtered PLP and 1 mL of 50 g/mL kanamycin. The resuspended cells were centrifuged again in a GS 3 rotor at 6000 rpm at 4 C for 40 minutes, to wash off any remaining rich media solution.

The supernatant was discarded, and the cells were resuspended in 500 mL of M9 minimal media supplemented with 3 g of glucomannan, 1 g chitin, 10 mL of a 50 % sterile filtered glucose solution, 50 g PLP and 500 L of 50 g/mL kanamycin. The cells were left shaking at 25 C at

250 rpm for 3 days. The cultures were then centrifuged in a GS 3 rotor at 6000 rpm at 4 C for

40 minutes, the supernatant was collected, purified and screened for glycosidase inhibition.

2.3.2.3 Purification methods

The cell culture supernatants were loaded onto an Amberlite IR 120 (H+) cation exchange column. The column had a diameter of 5 cm and was filled with resin until the 32 cm mark. The culture supernatant was loaded and washed with 1L of deionized water (dI H2O). The column

44

was then eluted with 0.5 M NH4OH and the alkaline fractions (pH ≥ 7.5) were collected and concentrated to dryness by rotary evaporation and analyzed for glycosidase inhibition. The dried residue was then extracted with a minimal amount of methanol (MeOH). The precipitate was separated from solution by filtration. The supernatant was then concentrated to dryness, resuspended in dI H2O and analyzed for maltase inhibition. The precipitate was also re-dissolved in dI H2O and analyzed for maltase inhibition to confirm it had no activity.

The supernatant, having shown maltase inhibition activity, was then loaded onto a Dowex

1x8 (Cl-) anion exchange column. The Dowex column had a 2.54 cm diameter and was filled with resin to the 32 cm mark. The column was eluted with dI H2O and 7 mL fractions were collected. Each fraction was then analyzed for maltase inhibition. Active fractions were collected, concentrated, resuspended in 1 mL of dI H2O and loaded the 1 mL onto the C18 HPLC

Phenomenex column (250 x 15 mm, 10 micron). The HPLC column was eluted with 1 % acetonitrile (ACN) at a rate of 3 mL/min and 6 mL fractions were collected. The fractions were analyzed for maltase inhibition, active fractions were collected, concentrated to dryness and re- dissolved in 0.5 mL dI H2O. The solution was loaded onto a Zorbax 300- SCX (9.4 x 250 mm, 5 micron) HPLC column. The column was eluted isocratically with 1 % 50 mM ammonium acetate (NH4Ac), pH 5 at a rate of 1 mL/min for 15 minutes and then ramped to 30 % 50 mM

NH4Ac, pH 5, over 5 minutes; 2 mL fractions were collected. The fractions were analyzed for maltase inhibition and the active fractions were concentrated down to dryness.

For removal of 4AD, the active fractions were re-dissolved in 0.5 mL NH4Ac, pH = 9, and loaded onto a polymer bound boronic acid resin that was prepped with 5 % acetic acid and rinsed with dI H2O. The column was 12.7 mm in diameter and loaded with 1 g of resin. After loading the sample, the column was washed with 3 x 3 mL of dI H2O with dropwise elution

45

followed by 1 % acetic acid elution. 1 mL fractions were collected and analyzed for maltase inhibition activity.

For the final purification step, the inhibitor mixture was dissolved in 0.5 mL of dI H2O and loaded onto a semi-prep PolyLC Polyhydroxyethyl A HILIC HPLC column. Solvent A was

100% ACN and solvent B was 6.5 mM NH4Ac, pH 5. Gradient elution was applied 0% B for 5 minutes, switched to 15% B with 5 min ramp time at a rate of 1 mL/min. Immediately after 15%

B was reached, the solvent was switched to 35% B with a 15 min ramp time. Again, immediately after 35% B was reached, the solvent was switched to 75% B with a 50 min ramp time. 2 mL

Fractions were collected with a fraction collector every 2 minutes. The fractions were then analyzed for maltase inhibition. Fractions that inhibited maltase were collected, concentrated down to dryness and then analyzed by 1H NMR and MS.

LC-MS experiments were carried out by the Mass Spectrometry Research and Education

Center at the University of Florida Chemistry Department. Samples were analyzed via LC-MS

(+) ESI. The mass spectrometer was an Agilent 6220 TOF with electro spray ionization (ESI) analyzed in the positive mode. Gas temperature was set to 350 º C, drying gas (N2)- 8.0 L/ min,

Nebulizer- 30 psig. HPLC used was an Agilent 1100 series system consisting of G13793 degasser and

G1312B binary pump. The column was a Thermo scientific, Accucore- 150-Amide-HILIC, 100 x 2.1.

Solvent A = 6.5 mM ammonium acetate in water, Solvent B= Acetonitrile 100%. Flow rate was 0.2 mL/min A:B= 10:90. The standard DAB-1 sample was dissolved in 2 mL methanol and the inhibitor extracted from C. pinensis as well as the deuterated inhibitor isolated from heterologous expression in E. coli were dissolved in 0.5 mL of methanol. The gradient was run as follows: from 0-5 minutes 95% B, from 5-14 minutes, gradient was switched from 95% B to 20% B, it was left at 20% B for 1 minute and then switched back to 95% B at 22 minutes and remained at 95% until minute 32.

46

2.3.3 Glycosidase Inhibition Assays

2.3.3.1  and -glucosidase assays

Each reaction mixture contained 1 mL of solution of 500 mM sodium phosphate buffer, pH 7.2, 50 U/mL of enzyme, and aliquots from purified fractions. The substrate concentrations were 50 M p-nitrophenyl--D-glucopyranoside (-PNP) for maltase (-glucosidase) assay and

400 M p-nitrophenyl--D-glucopyranoside (-PNP) for the -glucosidase. Control assays were run in parallel, containing all materials except aliquots from the purified fractions. The formation of p-nitrophenolate was monitored at 400 nm on a Shimadzu UV/Vis spectrophotometer for 5 minutes while maintaining the temperature at 25 C for yeast maltase and -glucosidase.

2.3.3.2 Assays on multi-well plate reader

A solution of 2 mg of yeast maltase in 500 µL of 500 mM sodium phosphate buffer, pH 7.2, was prepared (Solution A). In addition, a 10 mM -PNP in sodium phosphate buffer was prepared

(Solution B). For the assays, 60 µL of Solution A was added per 8 mL of buffer (Solution C) and

320 µL of Solution B was added per 8 mL of a separate buffer solution (Solution D). In a 96- well plate, the first well contained 180 µL of Solution C and the rest of the wells contained 160

µL of Solution C and 20 µL of each fraction from purification steps above. To initiate the reactions, 20 µL of Solution D was added to all the wells and were monitored for the release of p-nitrophenolate at 400 nm.

2.3.3.3 Jack Bean -mannosidase assay

This assay was developed as an endpoint assay because the enzyme functions at pH 5, precluding ready detection of p-nitrophenolate (pKa of 7.15). Because the protonated form of p-

47

nitrophenol is colorless, the reaction was quenched with a basic solution to monitor p- nitrophenolate at 400 nm. Each assay mixture contained purified fraction aliquots, 100 mM sodium acetate buffer at pH 5, 400 M p-nitrophenyl--D-mannopyranoside, and 58 U/mL of

-mannosidase in a total volume of 1 mL. The reactions were initiated at 37 C and then 300 L aliquots were removed at 3, 6, and 9-minute time points. These aliquots were then quenched with

0.1 M glycine buffer, pH 10.7 and measured at 400 nm.

48

CHAPTER 3 HETEROLOGOUS EXPRESSION OF AZASUGAR BIOSYNTHETIC SIGNATURE IN E. COLI

3.1 Overview

Bioinformatic analysis led us to a homologous azasugar signature three-gene cluster present within the C. pinensis genome. In Chapter 2 I presented evidence for C. pinensis’ production of a potent α-glucosidase inhibiting azasugar, DAB-1. However, is this homologous biosynthetic cluster present in C. pinensis’ genome responsible for the biosynthesis of DAB-1? In this chapter I report evidence on the elucidation of the function for the homologous azasugar biosynthetic cluster in C. pinensis. The homologous cluster (Cpin_2154, Cpin_2153, and Cpin_2152, NCBI reference sequence

NC_013132.1) was cloned into pETBlue-2. The resulting plasmid was used to transform in E. coli and resulted in the production of a slow-onset inhibitory azasugar, nectrisine.79

Reduction of nectrisine with NaBH4 or NaBD4 resulted in DAB-1 or DAB-1 deuterated at the anomeric carbon, which is supported by NMR and HRMS data.

3.2 Results and Discussion

For organisms that have been extensively studied and are easy to genetically manipulate, a classical method for learning about an enzyme’s function within its genome is to knock out the gene by disrupting its sequence. This was the method used by Clark et al 2011 to identify that GabT1 was an essential enzyme for the initiation of the biosynthesis of DNJ in B. amyloliquefaciens. Unfortunately, identifying the enzymes responsible for the biosynthesis of DAB-1 in C. pinensis is not as straight forward because there is yet to be any literature on procedures to genetically manipulate the C. pinensis species. There are some techniques for the genetic manipulation of Cytophaga-

49

Flavoobacterium-Bacteroides subphylum published.80–82 However , these techniques involve the use of a plasmids consisting of a transposon, which inserts itself randomly into the genome of the bacterium of interest. These methods of genetic manipulation have been mostly used to understand the mechanism(s) of bacterial gliding motility.80,83,84

Lacking a method to specifically target the putative azasugar biosynthetic cluster in C. pinensis, I attempted to genetically manipulate the C. pinensis genome by using published methods for gene knockout in E. coli and Bacillus.59,85 These attempts to knockout Cpin2154 (the putative aminotransferase) in C. pinensis were not successful.

Our next approach was to clone the putative azasugar cluster and express the genes in a well-studied organism such as E. coli. The putative azasugar biosynthetic cluster, Cpin2154, Cpin2153, and Cpin2152, consisting of a putative aminotransferase, alcohol dehydrogenase, and phosphatase were cloned into a pETBlue-2 vector. The resulting plasmid, termed pCluster, (Figure 3-1A) was transformed into E. coli Tuner cells, and protein expression was induced with IPTG at 25 ºC. The presence of three new protein bands after 1 day of IPTG induction corresponding to the predicted molecular weight of 45.9 kDa, 37.3 kDa, and 25.7 kDa for Cpin2154, Cpin2153, and Cpin2152, respectively, were observed by SDS- PAGE of crude cell lysates (Figure 3-1B). The appearance of these bands after induction suggested that the expression of the cluster in

E. coli was successful. Harvesting of E. coli cells after 1-day post-IPTG induction resulted in extracts exhibiting low maltase inhibition following cation exchange chromatography. Harvesting of the cells 3 days post-IPTG induction resulted in higher maltase inhibition activity after cation exchange purification. Therefore, after 3 days, the

E. coli cells were harvested by centrifugation. The cell pellets were resuspended in water

50

A) B) kDa T0 T1 MW T2 250 80 Putative 60 Aminotransferase 50 Cpin2154 40 Cpin2153 30 Cpin2152 25 Putative Dehydrogenase

Putative Phosphatase

Figure 3-1. A) pETBlue-CpinCluster plasmid used for heterologous expression in E. coli. B) SDS PAGE gel of pCluster expression in E. coli.*

*MW = molecular weight (MW) marker, T0 = before IPTG induction, T1 = 1 hr after induction, and T2 = 1 day after induction. The red dotted line links the MW marker band to the corresponding mass in kDA. Cpin2154, the GabT1 homolog, has a theoretical MW of 45.9 kDa and is marked by a purple arrow. Cpin2153, the GutB1 homolog, has a hypothetical MW of 37.3 kDa and is marked by a red arrow. Cpin2152 is the putative phosphatase, it has theoretical MW of 25.7 kDa and is marked by a green arrow on the protein gel image.

and sonicated to lyse the cells. Unlike the C. pinensis cell pellets, it was necessary to lyse

the E. coli cell pellets to maximize the yield of the inhibitor isolated. It was not necessary

to lyse the C. pinensis cells, possibly because C. pinensis contains the Cpin2151 (Figure

2-1), a putative major facilitator superfamily (MFS) transporter, which presumably

exports the inhibitor outside of the cell. The supernatant of the media and the supernatant

of the lysed cell pellets were loaded onto an Amberlite IR120 (H+) column and eluted

with water followed by 0.5 M NH4OH.

Before DAB-1 was identified as the azasugar produced by C. pinensis, it was

predicted that C. pinensis would make DNJ, due to its homologous azasugar biosynthetic

cluster. If it made DNJ, then like in B. amyloliquefaciens, the cluster should produce MJ,

which is a better inhibitor of α-mannosidase. Therefore, the E. coli extracts were

51

Maltase Inhibition Assay of Extracts from Cluster Expression in E. coli After Cation Exchange Column 100 80 60 40 20 0 48 50 53 54 56 57 58 59 62 63 64 65

Fractions % Inhibition Maltase Inhibition of %

No Incubation 5 min Incubation 15 min Incubation

Figure 3-2. Maltase assay inhibition profile of extracts from heterologous expression of the putative azasugar cluster in E. coli after Amberlite IR120 (H+) purification

analyzed first for α-mannosidase inhibition. I did not observe any obvious α-mannosidase inhibition from the E. coli extracts after cation exchange purification (Figure A-7); the slopes of the assays for the fractions were either the same or greater than the slope of the control. Next, the fractions were analyzed for inhibition against yeast maltase. I found that the E. coli pCluster extracts inhibited maltase and that the potency of the inhibition increased when the extract was pre-incubated with the enzyme for 15 minutes (Figure 3-

2).

To confirm that the inhibition and the slow-onset inhibition I was observing was real, the empty pETBlue-2 vector was expressed in E. coli as a control and did not show maltase inhibition (Figure A-8). This provides evidence for the inhibitor being produced as a result of expression of the enzymes in the cluster and not artifact due to a compound produced by E. coli. When comparing the kinetics of the maltase inhibitor isolated from heterologous expression in E. coli versus that of the inhibitor isolated from C. pinensis, the C. pinensis isolated inhibitor does not possess the same slow-tight binding; the

52

A) Maltase Inhibition Activity of Extract from E. coli Post Cation Exchange 0.1 0.08 0.06 0.04

0.02 Absorbamce 400 Absorbamce nm 0 1 30 59 88 117 146 175 204 233 262 291 Time (Seconds)

Control Maltase-Inhibitor 15 min Maltase-Inhibitor Pre-incubation

Maltase Inhibition Activity of Extract from C. pinensis Post Cation B) Exchange 0.1 0.08 0.06 0.04 0.02

Absorbance nm 400 Absorbance 0 1 30 59 88 117 146 175 204 233 262 291 Time (Seconds) Control Maltase-Inhibitor 30 min Maltase-Inhibitor Pre-incubation Figure 3-3. Comparison of time dependence maltase inhibition activity of extract from heterologous expression of putative azasugar cluster in E. coli (A) vs. extract from C. pinensis (B) post cation exchange purification potency of the inhibitor isolated from C. pinensis did not change over a 30 minute incubation period with the enzyme (Figure 3-3). This suggested that inhibitor being produced by the cluster in E. coli had a different structure than that of the C. pinensis isolated inhibitor. The E. coli isolated inhibitor also appeared to be less stable. This was inferred from the fact that the same purified sample exhibited weaker maltase inhibition a day after the IC50 was determined; a 3-fold decrease in inhibition was observed.

The slow-tight binding and unstable characteristics of the inhibitor isolated from

53

A)

Azasugar iminium-ion B)

Figure 3-4. A) Hypothesis for onset binding inhibition by inhibitor isolated from heterologous expression of putative azasugar cluster from C. pinensis in E. coli B) Reduction of iminium-moiety with NaBH4 heterologous expression of pCluster in E. coli led us to hypothesize that very much like in

B. amyloliquefaciens, the cluster in C. pinensis was making a precursor to DAB-1. We proposed that the instability was possibly due to a hydroxyl group on the anomeric carbon, which would make the molecule unstable because it can form an iminium species, known to form in some unstable azasugars,86 that can bind with a basic side chain within the glycosidase and covalently bind to the enzyme (Figure 3-4A). If this was the case, then the iminium-moiety could be reduced with NaBH4 (Figure 3-4B).

After the cation exchange column, a reduction step with NaBH4 was added to stabilize the compound. The extracts were analyzed for maltase inhibition and I observed that potent maltase inhibition was retained, but the inhibitor no longer exhibited slow- tight binding (Figure 3-5). Fractions consisting of the maltase inhibitor were collected and concentrated to dryness and further purified to identify the structure of the inhibitor.

The purification steps for isolation of the inhibitor were the same as the purification steps for the inhibitor isolated from C. pinensis, except for the lysis of the cell pellet and

NaBH4 reduction.

The IC50 after SCX HPLC was not measured because the mass of the inhibitor was less than 1 mg and therefore unable to accurately calculate the potency of the inhibitor. The 1H NMR spectrum of the reduced inhibitor produced by heterologous

54

Activity in Fractions from E. coli Post Cation Exchange and NaBH4 Reduction

0.1

0.08

0.06

0.04

PNP (µM) at 400 nm 400 at (µM) PNP 0.02

0 0 29 58 87 116 145 174 203 232 261 290

Time (Seconds) Control Maltase-Inhibitor Reduced_ Maltase-Inhibitor Reduced_15 min Maltase-Inhibitor Pre-incubation 15 min Maltase-Inhibitor Pre-incubation

Figure 3-5. Comparison of time dependence maltase inhibition activity of extract from heterologous expression of putative azasugar cluster in E. coli before and after NaBH4 reduction

Table 3-1. IC50 values for maltase by extracts from heterologous expression of pCluster in E. coli IC Purification 50 Dry Mass Weight from 20 L of Cultures (mg) (μg/mL) Cation Exchange Column * - NaBH4 reduction * 7700 MeOH Extraction 63 1250 Anion Exchange Column ≤2 40 C18-HPLC- ≤2 <1 SCX HPLC ND - *Individual fractions of the purifications were analyzed for maltase inhibition. The fractions were combined, but the concentrated potency of the combined fraction was not measured. expression of pCluster in E. coli after SCX HPLC purification strongly resembled 1H

NMR spectrum of the inhibitor isolated from C. pinensis cultures after SCX HPLC purification (Figure 3-6). The m/z 134 peak and the 152 in the MS chromatogram provides further evidence that, like in C. pinensis, I isolated DAB-1 and 4AD as a result of pCluster expression in E. coli (Figure 3-7).

55

I proposed that the homologous azasugar cluster in C. pinensis was responsible for the biosynthesis of nectrisine (23), an azasugar that possesses an imine functional

87 group and can be reduced to DAB-1 with NaBH4. If the cluster was biosynthesizing nectrisine, reduction with NaBD4 would lead to a deuterated DAB-1 (25) at the anomeric carbon, which would provide further evidence for the existence of the iminium moiety. In addition, reduction with NaBD4 would provide more insight on the mechanism of the conversion of nectrisine to DAB-1. From both cultures, C. pinensis and heterologous expression of the pCluster in E. coli, I extracted a mixture of DAB-1 (19) and 4AD (20).

Literature reports that 4AD has also been isolated from the fungus Thelonectria discophora and was identified as the substrate for the GutB1 homolog in the fungus.62 I hypothesized that, like in the Thelonectria discophora fungus, 4AD (20) oxidation is catalyzed by Cpin2153 (GutB1 homolog) to form (2S,3R,4R)-4-amino-2,3,5- trihydroxypentanal (21). Next, 21 spontaneously cyclizes to form (3S,4R,5R)-5-

(hydroxymethyl)pyrrolidine-2,3,4-triol (22), which can be further reduced to nectrisine

(23). If the equilibrium, post-Cpin2153 catalysis, leans to the left and 21 is accumulated, then reducing the culture extracts with NaBD4 should reveal a MS peak of 153.0941, corresponding to formation of deuterated 4AD (24). However, if the equilibrium leans to the right, post-Cpin2153 catalysis and 22 or nectrisine (23) is accumulated, then reduction with NaBD4 would lead to the production of deuterated DAB-1 (25) (Figure 3-

8). Reduction with NaBD4 would also help identify if the 4AD present formed as a result of reduction or if it is being made by the bacteria.

HRMS analysis of the extracts, from heterologous expression of the pCluster plasmid in E. coli, reduced with NaBD4 resulted in production of 4AD and deuterated

DAB-1; [M + H]+ peaks of 152.0921 and 135.0878, respectively, were observed (Figure

56

3-9). The [M + H]+ peak of 153.0941 was not observed, meaning that any 21 formed by oxidation of 4AD spontaneously cyclized to form either 22 or nectrisine. The presence of

4AD is most likely due to the inhibition of the Cpin2153 enzyme by some substance in the cell cultures, which reduced the rate of catalysis of 21 from 4AD.

The NaBD4 reduced extracts were still highly impure after SCX HPLC purification for 1H NMR analysis of the inhibitor (Figure C-3). As a result, the sample was further purified by HILIC HPLC. The fractions were assayed for maltase inhibition, and all active fractions were collected and then analyzed by 1H NMR, in which analysis of the chemical shifts led to the identification of deuterated DAB-1 (Figure 3-10). A 1H

COSY NMR is also available in the appendix Figure C-4. This strongly supports that an azasugar possessing an imine functional group, nectrisine, is produced by the azasugar biosynthetic cluster from C. pinensis. Nectrisine can be reduced by a putative reductase in C. pinensis, or by NaBH4 or NaBD4 in the case of heterologous expression of the cluster in E. coli, which does not produce an enzyme that can reduce nectrisine to DAB-

1. The biosynthesis of nectrisine by the cluster is further evidenced by the slow-onset inhibition of maltase, a characteristic reported in literature.88

In this chapter I provided evidence that the homologous azasugar biosynthetic cluster identified in C. pinensis initiates the biosynthesis of the azasugar DAB-1. The cluster synthesizes nectrisine, which can then be reduced by a putative reductase to DAB-

1. Nectrisine has only been isolated from plants and fungus, literature has not reported any sources for the ability of bacterial organisms to possess enzymes with ability to synthesize nectrisine.

57

A)

B)

Figure 3-6. A) 1H NMR (300 MHz) of inhibitor isolated from C. pinensis B) 1H NMR of reduced inhibitor isolated from E. coli post SCX HPLC in D2O

Figure 3-7. MS of NaBH4 reduced inhibitor isolated from heterologous expression of pCluster in E. coli post SCX HPLC

58

(20) (21) (22) (23)

+ [M + H]+ = 150.0722 [M + H]+ = 132.0616 [M + H]+ = 152.0878 [M + H] = 150.0722

(24) (25)

[M + H]+ = 135.0835 [M + H]+ = 153.0941

Figure 3-8. Scheme of possible products as a result of reduction of inhibitor with NaBD4 to elucidate the chemistry of the conversion of nectrisine to DAB-1

59

Figure 3-9. HRMS of NaBD4 reduced inhibitor isolated from heterologous expression of pCluster in E. coli post SCX HPLC

60

75% 25%

A)

5

3

2 4

1a 1b

B)

5 4 1a 2 3 1b

Figure 3-10. A) 1H NMR (300 MHz) of deuterated inhibitor isolated from E. coli after HILIC HPLC purification. B) 1H NMR (300 MHz) of DAB-1 standard.

61

3.3 Experimental

3.3.1 General Methods

The Escherichia coli Tuner pLacI (DE3) strain and the pETBlue-2 vector were obtained from Novagen. Oligonucleotides were purchased from IDT (Integrated DNA

Technologies). T4 DNA ligase, restriction enzymes, molecular weight standards, and

Phusion polymerase were purchased from New England BioLabs (NEB). Isopropyl-β-D- thiogalactopyranoside (IPTG) was purchased from Fisher Scientific. Sodium borohydride

(NaBH4) and sodium borodeuteride (NaBD4) were purchased from Sigma Aldrich. All constructs were confirmed by Sanger Sequencing core facility at the University of

Florida.

3.3.2 Cloning of the pETBlue2-Cpin2154-Cpin2153-Cpin2152 Expression Construct

The three gene cluster (Cpin2154, Cpin2153, and Cpin2152) was amplified from the C. pinensis strain DSM 2588 genomic DNA using 1 U of Phusion DNA polymerase by whole-cell PCR. For genomic DNA template, a patch of cells was streaked from Agar plates and added to 50 µL of dI H2O. The cells were denatured by heating at 98 °C for 5 minutes. The lysed cells were then centrifuged and 10 µL of the supernatant was added to the PCR reaction. Each PCR reaction mixture contained a final concentration of 1X of

5X Phusion HF Buffer, 200 µM of dNTPs, 0.5 µM Rev and Fwd Primers, and approximately 100 ng of template DNA. The samples were preheated at 98 °C for 1 minute and then proceeded with the following thermocycle: 98 °C for 30 s; then 30 x

(denaturation, 98 °C 10 s; annealing, 66 °C for 30 s; extension, 72 °C for 2 min) and a final extension at 72 °C for 10 minutes using the Fwd_gabt1_cluster_cpi and

Rev_yktc1_cluster_cpi primers (Table 3-2). The 2982 base pair PCR product was

62

digested with HindIII and XhoI restriction enzymes and then ligated into the HindIII/XhoI restriction sites of digested and purified pETBlue-2 vector. For restriction enzyme digest reactions, 1 unit of each restriction enzyme was added per 1 µg of DNA. Ligations were composed of 1 µL of T4 DNA Ligase Buffer (10X), a 1:3 ratio of vector DNA (100 ng): insert DNA, 1 uL of T4 DNA Ligase, and nuclease-free water for a 10 µL reaction. The ligation mixture (5 µL) was transformed into chemically competent E. coli TOP10 cells.

Miniprep DNA isolated from various transformants was screened by restriction analysis with BamHI. The plasmids (pCluster) showing the correct restricted bands were submitted for Sanger sequencing confirmation.

Table 3-2. List of deoxyoligonucleotides for expression of C. pinensis cluster and E. coli Primer Sequence (5’→3’) Restriction site Fwd_gabt1_cluster_cpi ATCGaagcttATGTCTGCTAACATTGAGTTCACAAAAGGC HindIII Rev_yktc1_cluster_cpi TCAGctcgagTCAGGAAGTTAATGTCAAAAGGCCAATATG XhoI

3.3.3 Expression of Cpin2154, Cpin2153, Cpin2152 in E. coli and purification of nectrisine

3.3.3.1 Rich media growth

The pETBlue-2 plasmid containing the C. pinensis cluster was co-transformed with the pRare plasmid in Tuner chemically competent cells and plated on LB agar plates supplemented with 34 g/mL chloramphenicol (to select for the pRare plasmid) and 50

g/mL of carbenicillin (to select for the pCluster plasmid) and incubated at 37 C overnight. Transformants were picked and grown in 5 mL LB media starter cultures at 37

C, shaking at 250 rpm, overnight. The starter cultures were then transferred to 500 mL

LB media cultures in 2 L baffled flasks, supplemented with 34 g/mL chloramphenicol

63

and 50 g/mL of carbenicillin at 37 C with shaking at 250 rpm. Once the OD600 reached

0.8, the cultures were induced with 500 M IPTG and incubated at 25 C with shaking at

250 rpm. After three days the cultures were harvested. As a control, E. coli cultures were transformed with empty pETBlue-2 vector and grown as described above.

3.3.3.2 Minimal media growth

E. coli cells were propagated in rich media conditions as described above. After overnight inoculation in LB broth, the 1 L cultures were centrifuged in a GS 3 rotor at

6000 rpm at 4 C for 40 minutes. The supernatant was discarded, and the cells were resuspended in 250 mL M9 minimal media (64 g Na2HPO4.7H2O, 15 g KH2PO4, 5 g

NH4Cl, 2.5 g NaCl, 2 mM MgSO4, 0.1mM CaCl2 per L at pH 7). The resuspended cells were centrifuged again in a GS 3 rotor at 6000 rpm at 4 C for 40 minutes, to wash off any remaining rich media solution. The supernatant was discarded, and the cells were resuspended in 500 mL of M9 minimal media supplemented with 10 mL of a 50 % sterile filtered glucose solution, 50 g PLP and 50 g mL-1 carbenicillin. The cultures were inoculated at 25 C while shaking at 250 rpm until an OD600 of 0.8 was reached. The cells were then induced with 500 M IPTG and incubated at 25 C with shaking at 250 rpm.

After 3 days the cultures were harvested, purified and screened for glycosidase inhibition.

3.3.3.3 Purification methods for compounds produced in E. coli

The purification methods were the same as described for C. pinensis in section

2.3.2 with the following differences: After three days, the E. coli cultures were harvested and centrifuged as described previously, the supernatant was loaded onto an Amberlite IR

120 (H+) column and instead of discarding the cell pellets, they were resuspended in 100

64

mL dI H2O and lysed by sonication on wet ice. The lysate was centrifuged for 30 minutes at 16,000 rpm at 4 °C. The supernatant of the centrifuged lysate was also loaded onto the

Amberlite column which was eluted with water followed by 0.5 M NH4OH. The alkaline fractions were collected, concentrated to dryness and treated with NaBH4 or NaBD4.

For NaBH4 or NaBD4 reduction, 400 mg of NaBH4 or NaBD4 was dissolved in 5 mL dI H2O. The concentrated Amberlite fractions containing inhibitor were re-dissolved in 40 mL of dI H2O, pH 9, and added to a 1 L round bottom flask with a stirring bar. The

5 mL solution of NaBH4 or NaBD4 was added to the round bottom flask dropwise while stirring. The solution was left stirring at room temperature, for approximately 12 hrs. The reaction was stopped by adding 1N H2SO4 until a pH 5 was reached. The solution was then concentrated to dryness, resuspended in 2 mL of dI H2O and approximately 200 mL

MeOH and concentrated to dryness. Resuspension in 2 mL of dI H2O and 200 mL MeOH was repeated 3 times to remove any excess borate from the solution. The reduced fraction was then re-dissolved in 50 mL of water (insoluble materials were removed by filtration) and analyzed for maltase inhibition.

The sample was concentrated to dryness again and further purified by MeOH extraction, anion exchange column, C18 HPLC, SCX HPLC, and polymer bound boronic acid resin as described for purification of C. pinensis inhibitor in section 2.3.2.

3.3.4 Glycosidase Inhibition Assays

Inhibition assays were carried out as described in 2.3.3.

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CHAPTER 4 BIOINFORMATIC ANALYSIS OF THE AZASUGAR BIOSYNTHETIC SIGNATURE

4.1 Overview

The isolation and characterization of natural products is a labor-intensive process involving fractionation, screening, and partial characterization of extracts, with chance playing a substantial factor on whether there will be a significant discovery or not.

Applying a combination of chemical analysis and bioinformatics analysis in natural products discovery helps to decrease the ‘chance’ component from the process. Despite the availability of computational aids in natural product discovery, the biosynthetic pathway of azasugars has only been partially identified, and despite their value as biologically active compounds, their ecological role remains unknown. Throughout the years, azasugar research has instead largely focused on their chemical synthesis and their potential application as pharmaceuticals and agrochemicals. With the increasing availability of whole genome sequences, I hope to identify new azasugar producing organisms and gain insight on how widespread azasugar production may be amongst different organisms. Identifying the organisms that produce azasugars and learning about the azasugar’s biosynthetic pathway could help elucidate the azasugar’s role in nature.

In this chapter I use bioinformatic tools to search existing genome sequences to find new organisms that may have the capability of biosynthesizing azasugars. More than

150 unique microorganisms containing a putative azasugar biosynthetic cluster were identified (Figure 4-1). The putative clusters include class III aminotransferases, haloacid dehalogenases which include some phosphatases, inositol phosphatase, and alcohol dehydrogenase (Figure 4-2).

66

Figure 4-1. Dendrogram of microorganisms containing a putative azasugar biosynthetic cluster.

(1) Aminotransferase: (7) (3) (1) Haloacid dehalogenase (1) (HAD, HAD_2): (1)

(2)

(1) Inositol Phosphatase: (2) (1) (25) Alcohol dehydrogenase (8) (ADH): (1) (1) (4) (1) Major Facilitator Family (1) Transporter (MFS): (1) (37) (31) Glycosyltransferase: (1) (1) (1) Oxidoreductase: (1) (1) (15) Glycosidase: (4) (1)

Figure 4-2. Genome neighborhood of microorganisms with putative azasugar biosynthetic cluster and other genes of interest. The numbers in parenthesis represent the total amount of strains within the species that were identified as putative azasugar producers. On the right, enzymes of interest are labeled and highlighted. In addition, MFS, Glycosyltransferases, and glycosidases, clustered with the three genes are included which may indicate biosynthesis of new uncharacterized azasugars. 67

4.2 Results and Discussion

Table 4-1. List of azasugar producers reported in literature

Azasugar Whole Genome Azasugar Organism Biosynthetic Cluster Sequence Availability Produced Presence Streptomyces roseochromogenes R-46833,56 NA ND NJ Streptomyces lavendulae SF-42533,56 NA ND NJ Streptomyces nojiriensis SF-42633,56 NA ND NJ Streptomyces subtrutilus ATCC 2746789 NA ND DNJ, DMJ Streptomyces lavendulae ATCC 3143457,90 NA ND DNJ, DMJ Streptomyces ficellus UC 543891 NA ND NJ Bacillus subtilis MORI 3k-8560 NA Y DNJ Bacillus atrophaeus ATCC 937220 NA ND DNJ Bacillus atrophaeus 194292 Y Y DNJ Bacillus amyloliquefaciens 140N92 NA Y DNJ Bacillus amyloliquefaciens FZB4222 Y Y DNJ Bacillus subtilis DSM70493 NA ND DNJ Bacillus amyloliquefaciens AS38519 NA ND DNJ Bacillus subtilis B419 NA ND DNJ Bacillus subtilis B294 NA ND DNJ Bacillus subtilis S1095 NA ND DNJ Paenibaillus polymyxa DSM 36596,97 Y Y DNJ Cylindrospermum licheniforme Kutzing ATCC NA ND DMDP 2941298 Cylindrospermum alatosporum F. E. Fritsch SAG NA ND DMDP 437998 Cylindrospermum uscicola Kutzing SAG 44.7998 NA ND DMDP Cylindrospermum stagnale Born. Et Flan SAG 45.7998 NA ND DMDP Cylindrospermum sp. 98 Y Y DMDP Thelonectria discophora SANK 1829287 NA Y Nectrisine NA = not available, ND = not determined, Y = yes sequence is available

Azasugars have been isolated from over 20 different microorganisms, including a

fungus. The genome sequences of only a few of these organisms have been reported in

literature. Technological improvements have facilitated and made the whole genome

sequencing process more affordable. Therefore, this list is expected to be enlarged as

more genome sequences become available. A list of known azasugar producing

microorganisms, genome and gene sequence availability, and the azasugar(s) produced

can be found in Table 4-1. As of now, all reported azasugar producers for which the

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genome sequence is available have a putative azasugar biosynthetic gene cluster. Of those organisms that are known azasugar producers and for which the cluster sequence is available, the only four organisms in which three-gene cluster is confirmed to be responsible for the biosynthesis of the azasugar: Bacillus subtilis MORI 3k-85, Bacillus amyloliquefaciens FZB4, Bacillus amyloliquefaciens 140N and Thelonectria discophora

SANK 18292. Bacillus amyloliquefaciens FZB4 is the only fully sequenced genome out of the four microorganisms; only the three-gene cluster is available for Bacillus subtilis

MORI 3k-85, Bacillus amyloliquefaciens 140N, and Thelonectria discophora SANK

18292.22,60,61,92

In Table 4-2, the putative azasugar biosynthetic clusters from organisms reported to be azasugar producers are compared to the azasugar biosynthetic cluster from B. amyloliquefaciens using blastx from the NCBI database. Blastx aligns translated nucleotide sequences against a protein database and gives a percent sequence coverage and percent identity score, which helps infer the function of the protein and a score of whether the proteins may be homologous or not. When looking at these data it is important to remember that a result that infers that proteins are not homologous does not necessarily mean that proteins do not have the same chemical function. Vice versa, proteins that have homology does not necessarily mean that they will have the same chemical function. However, as more gene sequences become available and protein functions are experimentally assigned, improved statistical computational methods with the capability of predicting enzymatic function of homologous, analogous, and non- homologous enzymes.99,100

Though not perfect, bioinformatics can be useful and has been shown to have success at identifying enzyme function by comparing gene sequences.101,102 In Table 4-2,

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Table 4-2. BLASTx statistics of putative azasugar biosynthetic genes of known azasugar producers compared to azasugar biosynthetic genes in B. amyloliquefaciens ylo Aminotran_3 HAD Zn_ADH % % % % % % Organism Coverage Identity Coverage Identity Coverage Identity Bacillus atrophaeus ATCC 9372 99 93 100 89 100 86 Bacillus atrophaeus 1942 99 99 100 89 100 86 Bacillus amyloliquefaciens 140N 99 99 99 96 99 99 Paenibaillus polymyxa DSM 365 96 48 29 53 91 32 Cylindrospermum sp. 96 26 17 28 53 26 Bacillus subtilis MORI 3k-85 100 92 97 86 100 81 Thelonectria discophora SANK 18292 93 24 35 36 22 43 Aminotran_3= aminotransferase class III family, HAD= haloacid dehalogenase family, Zn_ADH= zinc-binding alcohol dehydrogenase family.

it can be observed that all the Bacillus sp. have high protein sequence percent coverage and high percent identity in comparison to B. amyloli1quefaciens. Alignment score suggests that the proteins are nearly identical and therefore it is expected that the enzymes carry out the same chemistry. Experimental data has been reported that confirms the function of the three-gene cluster and the biosynthesis of DNJ in Bacillus subtilis

MORI 3k-85 and Bacillus amyloliquefaciens 140N.60,92 On the other hand, the function of the cluster has not been experimentally determined for the remaining Bacillus strains that have been reported to produce DNJ. The high homology of the three-gene cluster sequence and confirmed DNJ biosynthesis, however, strongly suggests that the identified three-gene cluster is responsible for DNJ biosynthesis.

By contrast, the aminotransferase and alcohol dehydrogenase from Paenibacillus polymyxa, which is also a known DNJ producer, are homologous to the aminotransferase and alcohol dehydrogenase from B. amyloliquefaciens, but the percentage identity is much lower. In addition, the putative phosphatase does not show any homology at all.

From these statistics we can deduce that to identify DNJ producers, having an aminotransferase and dehydrogenase homolog is enough. However, this is only one data point and more azasugar producers and gene cluster functionality need to be identified and experimentally confirmed to determine whether this is a common phenomenon or 70

not. In the case of the Cylindrospermum sp. bacteria and the fungus Thelonectria discophora, both producers of pyrrolidinic azasugars (DMDP and DAB-1 respectively), the only homologous enzyme based on sequence alignment is the class III aminotransferase. The putative phosphatase and alcohol dehydrogenase do not have any homology to the corresponding enzymes in B. amyloliquefaciens. The Cylindrospermum and Thelonectria statistical results are interesting for many reasons. For one they are two different organisms, a bacterium and a fungus, yet they both make five-membered rings azasugars. Second, this shows that the gene sequence from bacteria can be used to identify azasugar producers in fungi and vice-versa. Third, aminotransferase homology is sufficient for the identification of azasugar producers. Again, there is very little experimental data available and more work and analysis needs to be done to confirm whether this phenomenon is an outlier or a common between fungi and bacteria.

In Chapters 2 and 3 of this dissertation, I identify that C. pinensis also has an azasugar biosynthetic cluster and that it produces the pyrrolidine DAB-1, a potent α- glucosidase inhibitor. In Table 4-3, the putative azasugar biosynthetic clusters from the known azasugar producers are compared to the putative azasugar biosynthetic cluster from C. pinensis. The class III aminotransferase from C. pinensis is homologous to all the class III aminotransferases of the known azasugar producers; the enzyme has highest homology to the Paenibacillus polymyxa aminotransferase. The putative phosphatase from C. pinensis is not homologous to any of the haloacid dehalogenase protein families from the known azasugar producers, including Cylindrospermum sp. and Thelonectria discophora, which are also pyrrolidinic azasugar producers. The alcohol dehydrogenase enzyme from C. pinensis shares some homology with all of the alcohol dehydrogenases with the respective enzyme from the known azasugar producers except for the alcohol

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Table 4-3. BLASTx statistics of putative azasugar biosynthetic genes of known azasugar producers compared to azasugar biosynthetic genes in C.pin Aminotran_3 HAD Zn_ADH % % % % % % Organism Coverage Identity Coverage Identity Coverage Identity Bacillus atrophaeus ATCC 9372 99 47 3 71 99 32 Bacillus atrophaeus 1942 99 47 3 71 99 32 Bacillus amyloliquefaciens 140N 96 47 7 44 99 32 Paenibaillus polymyxa DSM 365 96 60 20 44 99 45 Cylindrospermum sp. 97 48 22 32 79 22 Bacillus subtilis MORI 3k-85 95 47 7 38 99 32 Thelonectria discophora SANK 18292 96 26 24 45 22 31 Bacillus amyloliquefaciens FZB42 95 47 5 38 97 32 Aminotran_3= aminotransferase class III family, HAD= haloacid dehalogenase family, Zn_ADH= zinc-binding alcohol dehydrogenase family.

dehydrogenase from Thelonectria discophora, even though they both synthesize the same azasugar. Thus, this shows that context is important too even though homology may not be the sole predictor of an azasugar biosynthetic cluster. What is common amongst all these confirmed azasugar producers is that they all have clustered enzymes that catalyze transamination, removal of phosphate esters, and oxidation of the liberated hydroxyl. The clustering of these enzymes may be more important than homology of the cluster.

All current known azasugar producers with genomic sequences available show homology between the aminotransferases and previous work in the Horenstein lab determined that the GabT1 enzyme is an essential enzyme from the azasugar biosynthetic cluster required for the biosynthesis of DNJ in B. amyloliquefaciens.22 For this reason,

GabT1 was used in the Enzyme Function Initiation Enzyme Similarity Tool (EFI-EST)103 to identify organisms specifically containing homologous class III aminotransferases.

Subsequently, the EFI Genome Neighborhood Network Tool (EFI-GNT) was used to condense similarity results into a total of 155 organisms which contain a cluster of sugar- binding enzymes with the desired putative functions: aminotransferase, phosphatase, and alcohol dehydrogenase (Figure 4-1 and 4-2). The EFI-EST database only provided homologous enzymes from bacterial sources. In Table 4-4, statistical data obtained from

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protein sequence alignments from organisms that are not reported in the literature as azasugar producers but contain the putative azasugar biosynthetic cluster are listed.

It can be observed from the statistical analysis that all of the class III aminotransferases from the organisms listed are homologous to GabT1 from B. amyloliquefaciens. This was to be expected as the aminotransferase was used to identify other possible unknown azasugar producers. The alcohol dehydrogenases from nearly all predicted azasugar producers have homology to the B. amyloliquefaciens’GabT1 enzyme, apart from a few organisms such as Propionibacterium acnes and some Streptomyces strains. The enzymes that are predicted to be phosphatases are the enzymes that show little to no homology to B. amyloliquefaciens’YktC1 enzyme. The fact that some of these organism’s phosphatases and alcohol dehydrogenases do not show homology does not take away the possibility that the may be azasugar producers, as can be seen in confirmed azasugar producers above. If future experimental data confirm that these organisms, in which all do not have homologous phosphatases or dehydrogenases, are azasugar producers, then this would confirm that sequence homology is not required, but the clustering of sugar binding enzymes with the functions of an aminotransferase, phosphatase, and alcohol dehydrogenase. No azasugar producers without the three-gene cluster have been identified. Therefore, based on current available data, it seems that the three-gene cluster, but not homology, is required for the biosynthesis of azasugars.

It is interesting to note that the majority of the organisms that appear to be potential azasugar producers are bacteria that have been isolated from the soil or marine sediment. Some hypotheses for microbial production of azasugars are that they help bacteria sequester food by inhibiting the glycosidases from other competing organisms or as a defense mechanism against other organisms by inhibiting their glycosidases. There is

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Table 4-4. BLASTx statistics of putative azasugar biosynthetic genes of unknown azasugar producerscompared to azasugar biosynthetic genes in B. amyloliquefaciens Aminotran_3 HAD Zn_ADH % % % % % % Organism Coverage Identity Coverage Identity Coverage Identity [Clostridium] populeti 89 42 9 37 93 28 [Flexibacter] sp. ATCC 35208 94 47 7 44 97 37 Acinetobacter pittii ANC 4050 88 37 None None 80 32 Actinobacteria bacterium OK074 96 60 95 56 92 39 Actinoplanes derwentensis 93 42 32 25 70 31 Actinoplanes philippinensis 92 42 11 28 74 33 Actinoplanes utahensis strain 91 41 26 26 64 32 NRRL12052 Aeromonas dhakensis strain F2S2- 93 35 19 25 74 30 1 Aeromonas sp RU34C 93 37 19 40 89 26 Bacillus amyloliquefaciens subsp. 99 100 100 99 100 99 plantarum UCMB 5033 Bacillus amyloliquefaciens 99 100 100 98 100 99 UMAF6639 Bacillus amyloliquefaciens Y2 99 100 100 99 100 99 Bacillus atrophaeus strain 99 92 100 89 100 86 B4144_201601 Bacillus cereus BAG1X1-1 98 84 99 82 98 74 Bacillus cereus BAG1X2-1 98 84 99 82 98 74 Bacillus cereus BAG1X2-2 98 84 99 82 98 74 Bacillus cereus BAG1X2-3 98 84 99 82 98 74 Bacillus cereus BAG2O-1 98 84 99 82 98 74 Bacillus cereus VD133 98 84 99 83 100 73 Bacillus cereus VD169 98 84 99 83 98 74 Bacillus gaemokensis strain 98 84 99 83 100 74 KCTC13318 Bacillus intestinalis strain 1731 99 92 100 86 100 80 Bacillus mycoides strain BTZ 98 84 99 82 100 74 plasmid pBTZ_1 Bacillus nakamurai strain NRRLB- 99 97 100 94 100 94 41092 Bacillus pseudomycoides DSM 94 84 99 82 100 74 12442 Bacillus pseudomycoides strain 98 84 99 82 100 74 AFS098564 Bacillus siamensis strain sdc15 99 99 100 98 100 99 Bacillus simplex strain SH-B26 98 83 99 82 98 74 Bacillus sp. 103 mf 98 83 99 82 100 73 Bacillus sp. 491mf 98 84 99 81 100 73 Bacillus sp. 5B6 99 99 100 99 100 99 Bacillus sp. FMQ74 99 92 100 86 100 80 Bacillus sp. JS 99 92 100 86 100 81 Bacillus sp. OK838 98 83 99 82 98 72 Bacillus sp. RUPDJ 99 100 100 99 100 99 Bacillus subtilis strain ATCC 99 99 100 97 100 99 13952 Bacillus subtilis subsp. 99 92 100 86 100 82 inaquosorum strain KCTC 13429 Bacillus subtilis subsp. spizizenii 99 92 100 86 100 79 TU-B-10 Bacillus thuringiensis serovar 98 84 99 83 98 74 aizawai strain T07151 Bacillus vallismortis strain NBIF- 99 99 100 99 100 99 001 Bacillus velezensis strain NKYL29 99 100 100 99 100 99 Bacillus velezensis strain V4 99 99 100 99 100 99 74

Table 4-4. Continued Aminotran_3 HAD Zn_ADH % % % % % % Organism Coverage Identity Coverage Identity Coverage Identity Chitinophaga filiformis 94 47 2 44 97 33 Chitinophaga pinensis DSM 2588 95 47 5 38 97 32 Chitinophaga rupis 94 47 27 71 97 32 Chitinophaga sp. CF118 94 48 12 64 97 33 Chitinophaga sp. YR573 94 47 7 35 97 34 Clostridium beijerinckii 92 44 None None 93 30 Clostridium gasigenes 93 42 20 31 93 32 Clostridium puniceum 92 44 11 33 93 32 Cosenzaea myxofaciens ATCC 88 40 None None 95 29 19692 Erwinia piriflorinigrans CFBP 93 37 None None 94 27 5888 Erwinia tasmaniensis strain 93 36 None None 96 28 ET1/99 Jeotgalibacillus marinus strain 99 92 100 86 100 79 DSM 1297 Kitasatospora aureofaciens strain 93 64 95 55 90 34 NRRLB-2658 Marininema mesophilum 98 80 95 75 100 70 Micromonospora rosaria strain 92 44 25 28 73 34 DSM 803 Mucilaginibacter lappiensis strain 95 47 34 71 97 33 ATCC BAA-1855 Mycobacterium abscessus subsp. 100 100 100 99 100 99 massiliense Myxococcus fulvus 124B02 93 47 None None 73 36 Paenibacillus amylolyticus strain 93 49 27 26 98 32 FSL 3-0122 Paenibacillus elgii strain M63 93 47 None None 98 34 Paenibacillus jamilae strain CN9 93 48 4 60 80 34 Paenibacillus lautus strain FSLF4- 93 49 None None 98 33 0100 Paenibacillus mucilaginosus 3016 94 48 None None 98 33 Paenibacillus mucilaginosus K02 94 48 None None 98 33 Paenibacillus mucilaginosus 94 48 None None 98 33 KNP414 Paenibacillus pabuli strain FSLF4- 93 49 15 29 98 33 0087 Paenibacillus peoriae strain 93 48 4 67 91 32 FSLA5-0030 Paenibacillus peoriae strain 93 48 4 67 50 45 FSLJ3-0120 Paenibacillus polymyxa E681 93 48 4 60 91 32 Paenibacillus polymyxa strain 93 48 4 67 91 32 DSM 365 Paenibacillus polysaccharolyticus 93 49 16 75 98 32 Paenibacillus rhizosphaerae strain 93 49 9 38 98 34 FSL R5-0378 Paenibacillus sp. A59 93 48 47 27 98 33 Paenibacillus sp. BD3526 93 48 6 27 91 31 Paenibacillus sp. CF095 93 49 27 27 98 32 Paenibacillus sp. D9 93 49 31 47 98 34 Paenibacillus sp. FSL R5-0765 93 48 27 26 98 32 Paenibacillus sp. FSL R5-192 93 49 27 27 98 32 Paenibacillus sp. IHB B 3084 93 48 18 47 90 32 Paenibacillus sp. MAEPY2 93 49 23 24 98 33 Paenibacillus sp. OK003 93 49 15 39 98 33 Paenibacillus sp. OK076 93 49 9 29 98 33 Paenibacillus sp. P22 93 49 31 47 98 34 Paenibacillus sp. SSG-1 93 49 9 38 98 34 Paenibacillus sp. TI45-13ar 93 49 42 26 98 32 75

Table 4-4. Continued Aminotran_3 HAD Zn_ADH % % % % % % Organism Coverage Identity Coverage Identity Coverage Identity Paenibacillus sp. VT-16-81 93 49 42 26 98 32 Paenibacillus tyrfis strain MSt1 93 47 14 39 98 35 Paenibacillus xylanexedens strain 93 49 27 27 98 32 PAMC 22703 Pedobacter cryoconitis strain 93 48 8 32 97 33 PAMC 27485 Pedobacter hartonius 93 47 4 100 97 34 Pedobacter lusitanus strain NL19 93 47 2 71 97 33 Photorhabdus asymbiotica subsp. asymbiotica (strain ATCC 91 37 5 35 87 28 43949/3105-77) Photorhabdus heterorhabditis 91 37 9 35 75 31 strain VMG Photorhabdus khanii NC19 93 36 5 35 80 28 Photorhabdus laumondii subsp. 91 37 5 35 75 31 laumondii strain TT01 Photorhabdus luminescens BA1 91 37 5 35 75 30 Photorhabdus luminescens strain 91 37 5 35 64 30 LN2 Photorhabdus temperata subsp. 91 37 5 35 75 31 Temperata Meg1 Photorhabdus temperata subsp. 91 37 5 35 75 31 thracensis strain DSM 15199 Propionibacterium acnes 96 58 2 86 13 30 HL201PA1 Saccharothrix sp. ALI-22-I 96 49 2 75 58 24 Serratia proteamaculans 88 39 9 32 95 27 Sinosporangium album 97 64 98 55 90 40 Streptococcus pneumoniae 99 99 100 97 100 99 Streptomyces albus subsp. albus 97 57 3 67 10 25 strain NRRLF-4371 Streptomyces autolyticus strain 97 57 3 67 10 25 CGMCC 0516 Streptomyces griseoflavus strain 96 63 95 54 90 33 NRRLB-1830 Streptomyces hygroscopicus subsp. 98 50 3 57 61 24 jinggangensis 5008 Streptomyces hygroscopicus subsp. 98 50 9 38 61 24 limoneus strain KCTC 1717 Streptomyces malaysiense strain 97 61 95 57 98 32 MUSC136 Streptomyces rimosus subsp. pseudoverticillatus strain NRRL 96 63 95 55 90 34 WC-3896 Streptomyces rimosus subsp. 93 64 95 55 90 34 rimosus ATCC 10970 Streptomyces rimosus subsp. 96 63 95 54 90 34 rimosus strain NRRL WC-3869 Streptomyces sp. 2314.4 92 58 95 58 96 35 Streptomyces sp. IMTB 2501 96 61 95 57 98 33 Streptomyces sp. NRRLF-5755 96 63 95 55 90 34 Streptomyces sp. PCS3-D2 isolate 97 61 95 57 97 32 P1 Streptomyces viridochromogenes 96 62 95 57 98 32 strain NRRL 3413 Vibrio neptunius strain S2394 94 39 7 30 50 37 Xenorhabdus beddingii 93 36 11 35 75 30 Xenorhabdus bovienii SS-2004 93 36 5 29 87 29 Xenorhabdus bovienii str. CS03 93 36 5 29 87 28 Xenorhabdus bovienii str. feltiae 93 36 5 29 87 28 Florida 76

Table 4-4. Continued Aminotran_3 HAD Zn_ADH % % % % % % Organism Coverage Identity Coverage Identity Coverage Identity Xenorhabdus bovienii str. feltiae 93 36 5 29 87 28 Moldova Xenorhabdus bovienii str. Kraussei 93 36 5 29 87 28 Becker Underwood Xenorhabdus bovienii str. Kraussei 93 36 5 29 87 28 Quebec Xenorhabdus bovienii str. 93 36 5 29 87 28 oregonense Xenorhabdus bovienii str. 93 36 5 29 87 28 Puntauvense Xenorhabdus cabanillasii JM26 93 35 5 29 75 30 Xenorhabdus doucetiae str. 93 36 6 38 90 29 FRM16 Xenorhabdus eapokensis strain 93 37 11 38 96 28 DL20 Xenorhabdus hominickii strain 93 36 16 50 80 30 DSM 17903 Xenorhabdus japonica 93 36 11 71 96 30 Xenorhabdus khoisanae strain 93 35 24 29 94 27 MCB Xenorhabdus mauleonii 93 35 11 29 76 29 Xenorhabdus nematophila AN6/1 93 37 35 71 90 28 Xenorhabdus nematophila ATCC 93 37 10 71 90 28 19061 Xenorhabdus nematophila F1 93 37 10 71 90 28 Xenorhabdus nematophila str. 93 37 35 71 90 28 websteri Xenorhabdus poinarii str. G6 93 36 5 29 95 28 Xenorhabdus sp. GDc328 93 37 11 38 90 28 Xenorhabdus thuongxuanensis 93 37 11 38 95 38 strain 30TX1 Xenorhabdus vietnamensis 93 36 9 35 96 28 Aminotran_3= aminotransferase class III family, HAD= haloacid dehalogenase family, Zn_ADH= zinc-binding alcohol dehydrogenase family.

also a possibility that the azasugars may play a role in the bacteria’s regulatory system by

inhibiting its own glycosidases. There is much work to be done in elucidating and

obtaining evidence on azasugars’ ecological function.

In order to be able to infer what the role of azasugars are, I would need to

understand more about the organisms which produce them. The following is a survey of

what is known of the possible azasugar producers listed in Table 4-4:

Clostridium. This genus of bacteria includes species that are known to inhabit

soils and the intestinal tract of animals, including humans. Some can be pathogenic and,

in humans, have been known to cause diarrhea.104

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Flexibacter. Genus includes species isolated from soil environments and have been identified as fish pathogens.105

Acinetobacter. Species from this genus have been found both in soil and water.

They are known to infect people with compromised immune systems in hospitals.106

Actinobacteria. Bacterial species from this genus have been found in both terrestrial and aquatic soil. They help decompose matter of dead organisms and are known to live symbiotically with plants.107

Aeromonas. The bacteria from this genus are mostly associated with human diseases. They can be found mostly in water sources.108

Bacillus. This genus is one of the most well studied and understood prokaryotes.

They have been isolated from soil bacteria and consist of both pathogenic and non- pathogenic strains. Many strains are used as biofertilizers due to their ability to help protect against fungus and insects. They have also been shown to promote plant growth.109,110 Some Bacillus sp. strains have been reported to be producers of azasugars such as DNJ and DMJ.

Chitinophaga. Not much is known about the species from this genus. They are found mostly in the soil and, as the name implies, are known to be able to hydrolyze chitin.69In this thesis I show evidence that Chitinophaga pinensis makes the azasugar

DAB-1.

Cosenzaea. This genus was re-classified and used to be known as Proteus. It consists of bacterial species that are normally part of the human and animal intestinal flora, but can be an opportunistic pathogen.111

Erwinia. Bacteria from this genus are mostly known to be plant pathogens that have been isolated from water, soil, seeds, and plant tissues.112

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Jeotalibacillus. All but one of the bacterial strains from this genus have been isolated from salty environments. The other was isolated from the soil and very little is known about these bacterial species.113

Kitasatospora. Bacteria belonging to this genus have been isolated from soil and sugar cane fields.114

Marininema. This genus is part of the Thermoactinomycetacceae family. They have been isolated from sea sediment, soil, and sugar cane fields.115

Mucilaginibacter. Not much is known about this genus except that bacterial strains have been isolated from soil and lichen.116

Mycobacterium. This genus includes bacterial species that are pathogenic to mammals. Some have been known to cause diseases in humans such as tuberculosis and leprosy. They live in water and food sources. Some have been found to be obligate parasites as they have never been found live outside of a host.117

Myxococcus. This genus consists of bacterial species isolated from Chinese sea water. It has be found to have lytic activity against bacteria and lower fungi.118

Paenibacillus. This genus was originally included within the Bacillus genus, but then reclassified as a separate genus. They have been isolated from many sources including humans, animals, and plants. These bacteria have played a significant role as biocontrol agents and as biofertilizers to promote plant growth.119 As mentioned previously in this chapter, the Paenibacilus polymyxa DSM 365 strain has been reported to produce the azasugar DNJ96.

Pedobacter. Bacteria belonging to this genus are mostly soil associated, but have also been recovered from water, chilled food, fish, and compost. They are a reservoir of antibiotic resistance genes.120

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Propionibacterium. This genus of bacteria can sometimes behave as parasites, but don’t necessarily need a host to complete its life cycle. They are known to live in symbiosis with humans and other animals. They usually do not cause problems for most people but can sometimes cause acne and other skin conditions. They can also be found in gastrointestinal tracts.121

Photorhabdus. Many bacteria from this genus are entomopathogenic and live symbiotically in a nematode’s gut. When the nematodes infect the insect, they release the bacteria into the insect’s blood and kills the insect within 48 hours.122

Saccharothrix. Bacteria from this genus have been isolated from soil. They have been found to produce beta-lysine-containing aminoglycoside antibiotics.123

Serratia. This genus is composed of opportunistic human pathogens. They tend to colonize the respiratory and urinary tracts. Bacteria sources include showers, toilet bowls, and wetted tiles. The Serratia proteamaculans strain, which I found to have a putative azasugar biosynthetic cluster, is a plant growth promoting bacteria.124

Sinosporangium. Bacteria from this genus have been isolated from soil.125

Streptococcus. This genus of bacteria is a normal inhabitant of the human respiratory track. When the immune system is compromised, some species can cause pneumonia and meningitis.126

Streptomyces. Many of the bacteria in this genus are isolated from soil. Most of the antibiotics that are currently being used have been isolated from Streptomyces strains.127 Glycosidase inhibitors such as valienamine, acarbose, and DNJ have been isolated from Streptomyces strains as well.21

Vibrio. Bacteria from this genus have been isolated from the marine environment and are known to cause food borne infections, usually from undercooked sea food.128

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Xenorhabdus. This genus of bacteria has been isolated from nematodes. Like in

Photorhabdus, they live symbiotically with the nematodes and are entomopathogenic. No free-living bacteria has ever been isolated outside of the nematode host. It has also been observed that the nematode cannot establish itself within the insect host if the bacteria is not present.122

In addition to many of the microorganisms predicted to be azasugar producers being mostly soil bacteria, from the brief survey on the microorganisms, many may live symbiotically with plants, nematodes, or even with humans. However, it is unknown if azasugars play a specific role in whether the microorganisms have a symbiotic or pathogenic relationship with other organisms.

Focusing specifically on the Photorhabdus and Xenorhabdus entomopathogenic capabilities, it is known that the nematodes cannot successfully infect the insect host without the bacteria, which is very fascinating. When the bacterium is present it is released into the insect’s blood, which is also interesting because azasugars (DNJ and

DAB-1 for example) are known for their potent inhibition of trehalase,129–132 an enzyme that catalyze the hydrolysis of the disaccharide trehalose. In addition, trehalose is the major blood sugar in insects and plays a crucial role as a source of energy and dealing with abiotic stresses.133 Therefore, I can infer that if these bacteria are actually azasugar producers, then azasugars might contribute to insect death via interference with trehalose metabolism. The bacteria that promote plant growth may be playing a similar role in which they release compounds such as azasugars to defend against other microorganisms and insects.

Additionally, the azasugar NJ, in particular, has been identified as having antibiotic properties against organisms such as Shigella, E. coli, S. lutea, and S.

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aureus.33,91,134 The role of antibiotics in nature are also being debated. Some hypotheses state that bacteria produce antibiotics to kill or inhibit competitors or as cooperative signals used for intercellular communication.135–137 Arguments against antibiotics as microbial weapons include that concentrations of antibiotics in the soil are too low to kill or inhibit bacteria.138 In addition, exposure of low concentrations of antibiotics may induce responses such as the formation of biofilm and expression of virulence genes, which may help the competing organisms rather than kill or inhibit it.139–142 These arguments have led the discussion against antibiotics as weapons of self-defense.

Nonetheless, experimental and computational analysis of antibiotic’s role in nature continue to show evidence that support the classic argument that their role is to mediate competition and not cooperative signals for the coordination of bacterial behaviors.137 As of now, other azasugars have not been identified as having antibiotic properties; either they do not exist, and this antibiotic property is only a characteristic of NJ in particular, or the particular strains they may have antibiotic resistance to have not been tested and identified.

Even if azasugars are not all antibiotics, many of the hypotheses for the role of antibiotics in nature can be applied to azasugars, only that they may have a different mode of action (for example, the targeting of glycosidases). Azasugars can act as weapons, weakening a competing or organism and eventually killing it, by inhibiting glycosidases. Like antibiotics, the inhibition of the adherence and formation of biofilms has been reported in the presence of DNJ.143 Exactly how DNJ inhibits biofilm formation is not yet well understood. Azasugars are also produced in low concentrations and their biosynthetic enzymes have been shown to have really slow turn overs.144

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In this chapter I have compared the protein sequences of the azasugar biosynthetic cluster of known azasugar producers and identify over 150 new microbial strains as possible azasugar producers. Considering the three enzymes we identify as a signature for azasugar biosynthesis, I found moderate homology across a wide range of species.

Overall, besides their potent inhibition of glycosidases and the antibiotic property of NJ, not much else is known about azasugars that could help deduce their role in nature.

Considerable work remains to do the wet biochemistry to isolate putative glycosidase inhibitors from these species and correlate their structures with the biosynthetic machinery identified via comparative genomics.

4.3 Experimental

The dendrogram of organisms with putative azasugar biosynthetic cluster was made by using the phyloT tree generator website (https://phylot.biobyte.de/ ) and providing the organisms NCBI taxonomy ID.

The Enzyme Function Initiation Enzyme Similarity Tool (EFI-EST)103 was used to identify organisms containing homologous GabT1 enzymes. Subsequently, the EFI

Genome Neighborhood Network Tool (EFI-GNT) was used to condense similarity results into organisms which contain a cluster of sugar-binding enzymes with the desired putative functions: aminotransferase, phosphatase, and alcohol dehydrogenase. Sequence alignments to determine % coverage and % identity was calculated using the Basic Local

Alignment Sequence Tool (BLAST) in the National Center for Biotechnology

Information’s (NCBI) website for the comparison of translated nucleotide to protein sequence (blastx).

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CHAPTER 5 CONCLUSIONS AND FUTURE DIRECTIONS

5.1 Conclusions

Using bioinformatics, I was able to use the gene sequences for the enzymes that are responsible for the biosynthesis of DNJ, to identify C. pinensis as a possible azasugar producing organism. NMR and HRMS data on extracts purified from the C. pinensis cultures led to the identification of the pyrrolidine azasugar, DAB-1. There is no literature on the bacterial production of DAB-1. The amino-alcohol, 4AD, was also isolated from the C. pinensis extracts.

The putative azasugar cluster was then cloned into a pETBlue-2 vector and expressed in E. coli to identify if the cluster is involved in the biosynthesis of DAB-1. The heterologous expression in E. coli led to the isolation of an unstable compound that exhibited slow-onset inhibition against the yeast maltase enzyme. Reduction of this inhibitor led to a more stable compound that retained the potent inhibition of yeast maltase and loss of slow-onset inhibition. HRMS and

NMR analysis of the extracts led to the identification of both DAB-1 and 4AD. Reduction of the inhibitor with NaBD4 led to the isolation of deuterated DAB-1, providing further evidence for the identity of the azasugar produced in E. coli as nectrisine.

I propose the following biosynthetic route for DAB-1 in C. pinensis based on the isolation of nectrisine, 4AD, and DAB-1 from the C. pinensis and E. coli extracts: glucose is converted to xylulose-5-phosphate (X5P) through the pentose phosphate pathway. X5P is aminated by Cpin2154 and dephosphorylated by Cpin2152 to produce 4AD. Cpin2153 then oxidizes 4AD, which undergoes spontaneous cyclization to form nectrisine. Finally, a putative reductase reduces nectrisine to produce the final pyrrolidine azasugar, DAB-1 (Figure 5-1).

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Figure 5-1. Scheme of proposed biosynthetic pathway for DAB-1 in C. pinensis

These results demonstrate that the existence of a homologous azasugar biosynthetic cluster within an organism is indicative of general azasugar production and not just one specific azasugar. Protein sequence alignments of the azasugar biosynthetic cluster from known azasugar producers show that all of the class III aminotransferases were homologous within organisms with genome sequences available. From these results, it appears that the class III aminotransferase may be the most important enzyme required for the identification of other possible azasugar producers. There may be certain advantages to the production of different azasugars amongst organisms that have this biosynthetic cluster. The azasugars that have been isolated thus far have been shown to have different selectivity profiles. Therefore, depending on the environment surrounding the azasugar producing organism, the azasugars may target specific glycosidases from other organisms to outcompete them for a food source or as a defense mechanism.

5.2 Future Directions

A lot of work remains to be done to fully understand the biosynthesis of azasugars. For example, though azasugars have been isolated from many sources and many applications for

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their use have been developed, the full biosynthetic pathway has not been identified. The three- gene cluster that has been identified only initiates the biosynthesis of DNJ, a possible epimerase and reductase that may be involved in the biosynthesis, have not yet been identified and do not appear to be clustered with the aminotransferase, phosphatase, and alcohol dehydrogenase.

Currently, the Horenstein group has been using bioinformatics to identify possible enzymes that may be responsible for the reduction of NJ to DNJ.

Knowledge on enzyme catalysis of azasugars is also limited by the lack of crystal structures. Though some enzymes within the azasugar biosynthetic cluster have been characterized, attempts at obtaining crystal structures for the enzymes have so far been unsuccessful. Crystal structures with substrate may be able to provide insight on the enzymes catalytic site and the essential amino acids involved in catalysis. With this information it may be possible to then use the sequence analysis combined with the essential catalytic residues to predict what specific azasugar an organism may produce.

Specifically related to the work in this dissertation, future projects could involve the isolation of each individual enzyme and their characterization. The feeding of carbon labeled carbohydrates could help elucidate the precursors for DAB-1 biosynthesis in C. pinensis, which can then be used for the characterization of the enzymes in the cluster. For the characterization of the GutB1 homolog, the 4AD substrate can be chemo-enzymatically synthesized by a D- fructose-6-phosphate aldolase (FSA) that has been reported as an enzyme with the capability of synthesizing a number of azasugars.145–148 This enzyme may also serve for the cheap biosynthesis of other azasugar precursors that can be fed to knockout strains of putative azasugar producers.

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In Figure 4-2, bioinformatic analysis revealed that there are other genes that may be of interest that appear to be clustered with the signature azasugar biosynthetic cluster. Many azasugar producers with the three-gene cluster have a putative MFS adjacent to the cluster, which may be involved in the exporting of the azasugar outside the cells or importing the required sugar substrates for the azasugar biosynthesis. Some also have glycosidases or glycosyltransferases clustered with the three genes that may be involved in the biosynthesis of possible azasugar polysaccharides.

For elucidation of the natural role of azasugars, it would be beneficial to confirm whether the organisms, with the three-gene cluster, identified in Chapter 4 are azasugar producers. Once azasugar production is confirmed, we can then knockout the GabT1 homolog—since GabT1 knockout in B. amyloliquefaciens was sufficient for removal of DNJ biosynthesis—in the microorganisms predicted to be azasugar producers. Microbial knockout is predicated on the ability to transform the bacteria, which has proven to be difficult for organisms that or not well studied. However, there are tools that have advanced our ability to genetically modify many organisms, such as the CRISPR-Cas9 system,149,150 and therefore knockouts for organisms that have no published methods of transformation may be possible.

Once microbial knockouts are obtained, experiments can be developed in which the phenotypes of the microbial knockouts can be compared to their wildtype in culture and in a controlled environment with possible symbionts and/or competitors. The bacterial genus

Xenoharbdus, may be the ideal bacteria to use for the analysis of the azasugar’s role in the bacteria-nematode symbiotic relationship. This is because there are published methods for the transformation and genetic manipulation of Xenoharbdus bacterial strains.151,152 Once the azasugar production is knocked out, we could look at how a knockout within the azasugar

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biosynthetic three-gene cluster affects the bacteria’s symbiotic relationship with the nematode. In addition, we can observe if the knockout increases or decreases the nematode’s ability to infect the insect, or if the insect has a higher chance of survival when infected by a nematode that hosts the mutated bacteria. These phenotypic differences could ultimately be used to propose evidence-based hypotheses regarding the azasugar’s natural role.

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APPENDIX A GLYCOSIDASE INHIBITION ASSAY FIGURES

Maltase Inhibition Assay of C. pinensis Extract After Cation Exchange 0.070 Column

0.060

0.050

0.040

0.030

0.020

0.010 Absorbance 400 nm Absorbance

0.000 0 50 100 150 200 Time (S) Control 0.136 mg Cpin extract 0.272 mg Cpin extract

Figure A-1. Yeast maltase assay of concentrated alkaline C. pinensis extract fractions from Amberlite IR120 (H+) purification. Adding 0.136 mg of extract resulted in 43 % inhibition enzyme, whereas 0.272 mg of extract gave 66 % inhibition

β-Glucosidase Inhibition Assay of C. pinensis Extracts After Cation 1.6 Exchange Column 1.4 1.2 1 0.8 0.6

0.4 Aborbance nm 400 Aborbance 0.2 0 4 6 8 10 12 14 16 Time (minutes) Control 7.3 mg Cpin extract

Figure A-2. Almonds β-glucosidase assay of concentrated alkaline C. pinensis extract fractions from Amberlite IR120 (H+) purification. Adding 7.3 mg of extract resulted in 15 % inhibition of the enzyme.

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Chitinase Inhibition Assay of C. pinensis Extract After Cation Exchange 0.45 Column

0.4

0.35

0.3

0.25

0.2 Absorbance nm) (560 Absorbance 0.15 0 10 20 30 40 50 60 70 Time (Minutes) Control 1.36 mg Cpin extract

Figure A-3. Streptomyces chitinase assay of concentrated alkaline C. pinensis extract fractions from Amberlite IR120 (H+) purification.

Adding 1.36 mg of extract resulted in 25 % inhibition of the enzyme. α-Mannosidase Inhibition Assay of C. pinensis Extract After Cation Exchange Column 1.6 1.4 1.2 1 0.8 0.6 0.4

Absorbance nm 400 Absorbance 0.2 0 0 2 4 6 8 10 12 14 16 Time (min) Control 0.7 mg Cpin extract

Figure A-4. Jack Bean mannosidase assay of concentrated alkaline C. pinensis extract fractions from Amberlite IR120 (H+) purification.

Adding 0.70 mg of extract resulted in 22 % inhibition of the enzyme.

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Maltase Inhibition Assay of Autoclaved vs Non-autoclaved C. pinensis Extract After Cation Exchange Column 0.045 0.040 0.035 0.030 0.025 0.020

Absorbance 400 nm Absorbance 0.015 0.010 0.005 0.000 0 50 100 150 200 250 300 Time (min) Control 199 mg autoclaved Cpin extract 395 mg non-autoclaved Cpin extract

Figure A-5. Yeast maltase assay of concentrated alkaline autoclaved and non-autoclaved C. pinensis extract fractions from Amberlite IR120 (H+) purification.

Addition of 199 mg of autoclaved extract to reaction mixture resulted in 33 % inhibition of the enzyme. Whereas, 395 mg of non-autoclaved extract resulted in 36 % of the enzyme. Maltase Inhibition Assay of ATCC 1565 Media After Cation Exchange Column 0.120 0.100 0.080 0.060 0.040

0.020 Absorbance Absorbance 400 nm 0.000 0 50 100 150 200 250 300 Time (min) Control 526 mg media extract 1052 mg media extract

Figure A-6. Yeast maltase assay of concentrated alkaline ATCC 1565 media fractions from Amberlite IR120 (H+) purification.

Addition of 1052 mg of extract to reaction mixture resulted in no inhibition of the enzyme.

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α-Mannosidase Inhibition Assay of Extracts from Cluster Expression in control 0.2 E. coli F45 0.18 F47 F49 0.16 F50 0.14 F52 F54 0.12 F55 F56 0.1 F57 0.08 F60 F63 0.06 F65 Absorbance 400 nm Absorbance 0.04 F66 F67 0.02 F68 0 F69 F70 3 4 5 6 7 8 9 Time (minutes)

Figure A-7. Jack Bean mannosidase assay of alkaline E. coli pCluster extract fractions from Amberlite IR120 (H+) purification.

Fractions did not show any apparent inhibition of the mannosidase, providing evidence for an alternative azasugar being produced by the cluster in C. pinensis in contrast to mannojirimycin produced by the cluster in B. amyloliquefaciens

Maltase Inhibition Assay of Fractions Post Cation Purification of E. coli pETBlue-2 100 80 60 40 20

Inhibtion % Inhibtion 0 -20 -40 1 3 5 7 9 11 13 15 17 19 21 23 25 27 29 31 33 35 37 39 41 43 45 47 Fraction # Figure A-8. Yeast maltase inhibition assay of alkaline E. coli pETBlue-2 extract fractions from Amberlite IR120 (H+) purification.

Alkaline fractions from expression of empty pETBlue-2 vector in E. coli did not show inhibition against maltase, providing evidence that inhibitor isolated from heterologous expression of the cluster in E. coli is due to the expression of the three genes.

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APPENDIX B HPLC AND ELSD DATA

A) HILIC HPLC Chromatogram 1.6 1.4 1.2 1 0.8

0.6 Absorbance 0.4 0.2 0 20 40 60 80 100 120

F1 F2 F3 F4 F5 Time (minutes) ELSD UV (220 nm)

B) Maltase Inhibitioon Assay of Fractions Post HILIC HPLC of C. pinensis Extract 100 80 60 40 20 0

% Inhibition Maltase Inhibition of % 1 2 3 4 5 Fraction #

Figure B-1. A) Chromatogram of C. pinensis extract after HILIC HPLC. B) Maltase inhibition profile of corresponding HILIC HPLC fractions.

This HILIC run was an initial attempt at purification of the inhibitor produced by C. pinensis using a gradient of water and methanol. As can be seen from the chromatogram, using methanol as the solvent on the HILIC column was not successful in removing all of the impurities.

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APPENDIX C NMR DATA

A)

B)

Figure C-1. A) 1H NMR (500 MHz) corresponding to maltase inhibitor from C. pinensis after HILIC HPLC purification. B) 1H NMR (500 MHz) of standard D/L N-acetylarigine.

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Figure C-2. 1H NMR (300 MHz) corresponding to maltase inhibitor from C. pinensis after HILIC HPLC.

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1 Figure C-3. H NMR (300 MHz) corresponding to maltase inhibitor from NaBD4 reduction of inhibitor extracted from E.coi heterologous expression after SCX HPLC purification.

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5

3 2 4 1a 1b

1 Figure C-4. H COSY NMR (300 MHz) of NaBD4 reduced extracts, from E. coli heterologous expression, after HILIC HPLC purification

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APPENDIX D HRMS AND MS DATA

Figure D-1. HRMS of C. pinensis extract after HILIC HPLC Purification.

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Figure D-2. HRMS of D/L- N-acetylarginine standard.

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Figure D-3. HRMS of C. pinensis extract from cultures supplemented with chitin, sorbitol, and glucose after HILIC HPLC.

100

152.0916

Figure D-4. HRMS of C. pinensis extract from cultures supplemented with chitin, glucose, and glucomannan after SCX HPLC.

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APPENDIX E NUCLEIC ACID AND AMINO ACID SEQUENCES

Figure E-1. Cpin2154 gene sequence.

Figure E-2. Cpin2154 amino acid sequence.

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Figure E-3. Cpin2153 gene sequence.

Figure E-4. Cpin2153 amino acid sequence.

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Figure E-5. Cpin2152 gene sequence.

Figure E-6. Cpin2152 amino acid sequence.

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APPENDIX F AGAR PLATES AND C. PINENSIS CULTURES

A) B)

Greenish-gray solid

C. pinensis culture

Chitinophaga pinensis cultures, August 10, 2018. Courtesy of Claribel Nuñez

Figure F-1. A) On the left, a 500 mL culture of C. pinensis in ATCC 1565 media supplemented with chitin, glucose, and glucomannan after 3 days. On the right, a 50 mL culture of C. pinensis in same conditions as the large culture n the left. Both have 50 µg/mL kanamycin. B) zoomed in picture of the 500 mL culture in A.

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B) 3)

1) C) 2)

A) 2) 3)

1) 1) 3) 2)

2)

3) 1) D)

Chitinophaga pinensis agar plates, August 13, 2018. Courtesy of Claribel Nuñez Figure F-2. ATCC 1565 agar plates of C. pinensis culture streaks from Figure F-1B.

D) 3)

1) A) 1)

2) 3)

2) C) 2) B) 3) 3) 1)

1)

2) Chitinophaga pinensis agar plates, August 13, 2018. Courtesy of Claribel Nuñez Figure F-3. ATCC 1565 agar plates of greenish gray solid from C. pinensis cultures from Figure F-1A. *Pictures of plates were taken after 3 days of growth at 25 ºC. A) Plate with no antibiotic, B) plate with 10 µg/mL neomycin, C) plate with 50 µg/mL of carbenicillin, and D) plate with 50 µg/mL kanamycin. The plates are divided into three sections. 1) is the first streak from the culture, 2) is the first dilution streak from 1, and 3) is the second dilution streak from 2.

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C) D) 2)

3)

2)

1)

3) A)

3) 1) 1) 3) 2) B) 2)

1)

Chitinophaga pinensis agar plates , August 13, 2018. Courtesy of Claribel Nuñez Figure F-4. ATCC 1565 agar plates of C. pinensis from frozen stock.

*Pictures of plates were taken after 3 days of growth at 25 ºC. A) Plate with no antibiotic, B) plate with 10 µg/mL neomycin, C) plate with 50 µg/mL of carbenicillin, and D) plate with 50 µg/mL kanamycin. The plates are divided into three sections. 1) is the first streak from the culture, 2) is the first dilution streak from 1, and 3) is the second dilution streak from 2

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APPENDIX G TIC AND EIC DATA

Standard DAB-1

Deuterated DAB-1

DAB-1 from C. pinensis

Figure G-1. TIC of standard DAB-1, deuterated DAB-1 isolated from heterologous expression of azasugar biosynthetic cluster in E. coli, and DAB-1 isolated from C. pinensis

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Standard DAB-1

Deuterated DAB-1

DAB-1 from C. pinensis

Figure G-2. EIC of m/z 134 of standard DAB-1, deuterated DAB-1 isolated from heterologous expression of azasugar biosynthetic cluster in E. coli, and DAB-1 isolated from C. pinensis

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Standard DAB-1

Deuterated DAB-1

Figure G-3. EIC of m/z 135 of standard DAB-1 and deuterated DAB-1 isolated from heterologous expression of azasugar biosynthetic cluster in E. coli

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BIOGRAPHICAL SKETCH

Claribel Nuñez was born in New York City. She grew up in Bushwick and Crown

Heights in Brooklyn, NY as the second oldest of five children. In 2010 she was the first in her family to graduate with a Bachelor of Science in chemistry from the City University of New

York at Brooklyn College. As an undergraduate she joined the MARC program and was trained as an organic chemist in Maria Contel’s lab. She then joined the Bridge to the PhD program at

Columbia University from 2010-2012 and was trained as a biochemist in Ann McDermott’s laboratory. In the summer of 2013 she joined Professor Horenstein’s research group at the

University of Florida to pursue a PhD in chemistry, specifically in the biochemistry division. She completed her PhD in May 2019.

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