The Pennsylvania State University

The Graduate School

Department of Biochemistry and Molecular Biology

ELUCIDATION OF THE DEVELOPMENTAL AND PHYSIOLOGICAL

ROLES OF NAD+ BIOSYNTHETIC PATHWAYS

A Dissertation in

Biochemistry, Microbiology and Molecular Biology

by

Melanie R. McReynolds

© 2017 Melanie R. McReynolds

Submitted in Partial Fulfillment of the Requirements for the Degree of

Doctor of Philosophy

August 2017

The dissertation of Melanie R. McReynolds was reviewed and approved* by the following:

Wendy Hanna-Rose Associate Professor of Biochemistry and Molecular Biology Dissertation Advisor Chair of Committee

Craig E. Cameron Professor of Biochemistry and Molecular Biology Eberly Chair in Biochemistry and Molecular Biology

Teh-hui Kao Distinguished Professor of Biochemistry and Molecular Biology Chair of Plant Biology Graduate Program

Lorraine Santy Associate Professor of Biochemistry and Molecular Biology

Pamela A. Hankey-Giblin Professor of Immunology

Andrew D. Patterson Associate Professor of Molecular Toxicology Special Signatory

David Gilmour Professor of Molecular and Cell Biology Co-Director for Graduate Education in BMMB

*Signatures are on file in the Graduate School

iii

ABSTRACT

NAD+ biosynthesis has proven to be an attractive and promising therapeutic target for influencing healthspan and obesity-related phenotypes as well as tumor growth. However,

NAD+ is a key metabolite that impacts the entire metabolome. Therefore, it is necessary to elucidate exactly how manipulating NAD+ biosynthetic pathways can lead to therapeutic benefits. Also, it is imperative to characterize the unexpected adverse reactions to manipulating the biosynthetic pathways to fully utilize this target for drug discovery. The goal of our research is to understand how NAD+ homeostasis is maintained to support its core metabolic roles and its signaling and regulatory roles involving NAD+ consumers. In this work, I investigate the developmental and physiological role of NAD+ biosynthetic pathways in C. elegans, their homeostatic interactions, and I reveal a biosynthetic pathway involving an enzyme outside of

NAD+ biosynthesis.

NAD+ is synthesized via distinct routes including de novo synthesis from tryptophan, salvage synthesis from nicotinamide, which feeds into the Preiss-Handler pathway from nicotinic acid in C. elegans, and via the phosphorylation of nicotinamide ribosides or nicotinic acid ribosides using nicotinamide riboside kinase (NMRK). We previously discovered that NAD+ salvage synthesis through the nicotinamidase PNC-1 is required for normal progression of gonad development in C. elegans. Global metabolic profiling suggested that glycolysis was perturbed in our pnc-1 mutants, which have lower global levels of NAD+. Furthermore, we were able to link compromised glycolysis to gonad delay in our loss of salvage NAD+ synthesis mutants. I investigated this model and demonstrated using metabolic carbon tracing that glycolysis is compromised in our pnc-1 mutants. iv

It’s been reported in the literature that C. elegans lack the de novo NAD+ biosynthetic pathway because quinolinic acid phosphoribosyltransferase (QPRTase) is not encoded in the genome. However, all genes coding for the key enzymes required for production of quinolinic acid (QA) from tryptophan are present in the C. elegans genome. Using metabolic deuterium tracing I revealed that de novo NAD+ synthesis from tryptophan is active. I also demonstrated that UMPS-1 as the enzyme responsible required for converting QA into NAD+ during de novo biosynthesis. In addition to this, I discovered a novel role for NMRK-mediated synthesis in embryonic hatching in C. elegans. Finally, I uncovered a compensatory network amongst the biosynthetic pathways that maintains NAD+ homeostasis.

In summary, this work has expanded our knowledge of the developmental and physiological roles of NAD+ biosynthetic pathways. Metabolic carbon tracing was implemented as a tool to examine metabolic flux in C. elegans. Also, this work suggests that an underground metabolic mechanism may contribute to NAD+ biosynthesis. The conserved enzyme UMPS-1 is substituting for the missing QPRTase, raising questions about the relevance of similar underground metabolic activity in higher organisms. This work associates a novel C. elegans’ hatching phenotype to NAD+ biosynthesis. Finally, this work deciphers the impact of manipulating NAD+ biosynthesis for therapeutics.

v

TABLE OF CONTENTS

List of Figures………………………………………………………………………………...…viii

List of Tables…………………………………………………………...………………...………xi

List of Abbreviations…………………………………………………………………………….xii

Acknowledgments………………………………………………………………………………xiv

Chapter 1: Introduction……………………………………………………………………………1

Part I: NAD+ is a central hub in cellular metabolism……………………………………..1 + Historical context of Vitamin B3 as a precursor for NAD ……………………….1 Role of NAD+ in redox reactions …………………………………………………2 Role of NAD+ as a substrate ……………………………………………………...3 Part II: Eukaryotic NAD+ biosynthesis…………………………………………………... 4 Overview of NAD+ biosynthesis………………………………………………… 4 Salvage NAD+ synthesis…………………………………………………. 4 Preiss-Handler Pathway………………………………………………….. 5 de novo NAD+ synthesis…………………………………………………..5 Riboside synthesis………………………………………………………... 6 Part III: Targets for NAD+ biosynthesis and metabolism based drug discovery………….8 Cancer inhibition……………………………………………………………..……8 Health and lifespan benefits.……………………………………………………..10 Neurological disorders…………………………………………………………...11 Novel antibiotics………………………………………………………………... 12 Part IV: Role of NAD+ biosynthesis in metabolic homeostasis………………………... 13

Chapter 2: Metabolic carbon tracing reveals disrupted glycolysis due to a loss of salvage NAD+ synthesis in C. elegans………………………………………………………………….. 15

Introduction………………………………………………………………………………15 Importance of NAD+ in energy-producing redox reactions……………………...15 Relationship between glucose metabolism and cellular NAD+ pools…………... 16 Loss of NAD+ salvage synthesis disrupts glycolysis leading to developmental reproductive delay in C. elegans………………………………………………... 17 Results…………………………………………………………………………………... 19 Development of metabolic tracing protocol to model compromised glycolysis in pnc-1 mutants………………………………………………………………… 19 Metabolic tracing supports compromised glycolysis in pnc-1 mutants………… 20 Glucose storage in pnc-1 mutants………………………………………………. 24 Discussion………………………………………………………………………………. 26 Compromised NAD+ biosynthesis leads to disrupted glycolysis………………. 26 Requirement to maintain a supply of carbon for oxidative phosphorylation by the mitochondria…………………..……………………………………………. 27 vi

Disruption of NAD+ metabolism leading to developmental delay can model tumor growth and progression…………………………...... …………. 29 Materials and Methods…………………………………………………………………...31 C. elegans Culture and Strains…………………………………………………...31 Targeted Metabolomics………………………………………………………….31 Metabolic Tracing with Stable Isotopes………………………………………....32 Quantitative Real-Time Polymerase Chain Reaction (qRT-PCR)……………….32

Chapter 3:Eukaryotic de novo NAD+ biosynthesis from tryptophan in the absence of a QPRTase homolog………………………………………………………………………………… 34

Introduction……………………………………………………………………………... 34 Results…………………………………………………………………………………... 37 NAD+ de novo synthesis contributes to NAD+ biosynthetic capacity………….. 37 Supplementation with NAD+ de novo precursors reverses NAD+-dependent phenotypes……………………………………………………………………….40 UMPS-1 is required for QA label to be incorporated into NAD+ biosynthesis….43 Loss of kyneurine pathway affects reproductive development…………………..47 Discussion………………………………………………………………………………..48 Intact de novo NAD+ biosynthesis in the absence of QPRTase homolog……….48 Requirement for NAD+ de novo biosynthesis for normal reproduction….……. 49 Materials and Methods………………………………………………………………….. 50 C. elegans Culture and Strains…………………………………………………. 50 Metabolite Supplementation……………………………………………………. 50 Phenotypic Analysis…………………………………………………………….. 51 Targeted Metabolomics………………………………………………………… 51 Metabolic Tracing with Stable Isotopes………………………………………... 52 Quantitative Real-Time Polymerase Chain Reaction (qRT-PCR)……………… 53

Chapter 4: Nicotinamide Riboside contributes to NAD+ biosynthesis and embryonic hatching in C. elegans……………………………………………………………………………..….54 Introduction……………………………………………………………………………... 54 Nicotinamide riboside as a precursor for NAD+ biosynthesis…………..……… 54 Results…………………………………………………………………………………... 56 Supplementation with NR reverses NAD+-dependent phenotypes……………...56 NR contributes to NAD+ biosynthesis………………………………………….. 58 NR contributes to embryonic hatching during development………………….... 60 Discussion………………………………………………………………………………. 63 NR contribution to the cellular NAD+ pool……………………………………...63 NR contribution to C. elegans’ embryogenesis………………………………….64 Materials and Methods…………………………………………………………………...66 C. elegans strains and culture……………………………………………………66 Metabolite supplementation…………………………………………….………..66 Targeted metabolomics…………………………………………………………..67 Phenotypic Analysis……………………………………………………………...68 Quantitative Real-Time Polymerase Chain Reaction (qRT-PCR)……………….68 vii

Chapter 5: Compensatory roles for NAD+ biosynthetic pathways and consumers in C. elegans………………………………………………………………………………..70 Introduction………………………………………………………………………………70 Critical nature of NAD+ pool in cellular metabolism……………………………70 Results……………………………………………………………………………………72 NAD/NADH ratio is not impacted in loss of salvage NAD+ synthesis mutants...... 72 PARP deletion increases NAD+ levels in loss of salvage NAD+ synthesis mutants…………………………………………………………………...73 Loss of NAD+ biosynthetic pathways results in a homeostatic response………..74 Loss of NAD+ biosynthetic pathways results in global metabolic changes……..75 Mitochondria are protected in NAD+ biosynthetic mutants……………………..82 Discussion……………………………………………………………………………….83 Compensatory network within NAD+ biosynthetic pathways and consumers to maintain global NAD+ homeostasis……………………………………...83 Material and Methods……………………………………………………………………86 C. elegans Culture and Strains…………………………………………………...86 NAD/NADH ratio………………………………………………………………..86 Targeted Metabolomics………………………………………………………….86 Phenotypic Analysis……………………………………………………………..87 Quantitative Real-Time Polymerase Chain Reaction (qRT-PCR)………………88

Chapter 6: Discussion……………………………………………………………………………89 Developmental and physiological roles of NAD+ biosynthetic pathways……………….89 Metabolic tracing reveals compromised glycolysis in pnc-1 mutants…………...89 de novo NAD+ synthesis from tryptophan in C. elegans……………………………91 NMRK-mediated synthesis in C. elegans………………………………………..93 Homeostatic interactions amongst NAD+ biosynthetic pathways in C. elegans...94 Compensatory network to maintain NAD+ homeostasis………………………………...95 Unique and diverse biological functions for NAD+ biosynthetic pathways involved in C. elegans reproductive development……………………………………………97 Underground metabolism: Alternative routes to synthesize NAD+……………………...98

References………………………………………………………………………………100 Appendix: Amino acids are not used as an energy source in pnc-1 mutants…………...110

viii

LIST OF FIGURES

Figure 2-1: NAD+ biosynthesis in C. elegans…………………………………………...... 7

Figure 1-3: Therapeutic targets for NAD+ biosynthesis and metabolism………………………. 13

Figure 2-1: Schematic of disrupted glycolysis in pnc-1(pK9605) mutants…………………….. 18

Figure 2-2: Scheme for metabolic carbon tracing in C. elegans………………………………... 19

Figure 2-3: Representative raw data for glucose and isotopic glucose peaks………...………….21

Figure 2-4. Fraction of total glucose pool labeled decreases in pnc-1 mutants……………….... 22

Figure 2-5: Glycolysis is blocked at the NAD+ dependent step in pnc-1 mutants……………... 22

Figure 2-6: Flow rate of isotopic label after pyruvate into lactate is equal to N2 in pnc-1

mutants………………………………………………………………………………….. 23

Figure 2-7: Flow of isotopic label after pyruvate into the TCA cycle in pnc-1 mutants……….. 24

Figure 2-8: Glucose transporter activity is up regulated in pnc-1 mutants……………...... 25

Figure 2-9: Trehalose steady state levels are increased, but fraction of total trehalose pool

labeled decreases in pnc-1 mutants……………………………………………...... 26

Figure 2-10: Compromised glycolysis due to disruption in NAD+ salvage synthesis models

tumor growth and progression…………………………………………..……………...... 29

Figure 3-1: Schematic of NAD+ de novo synthesis in C. elegans……………………………... 36

Figure 3-2: Loss of kynu-1 decreases global NAD+ levels…………………………………….. 37

Figure 3-3: Trp pool is isotopically labeled in N2 and kynu-1 and label from Trp into QA

is undetectable in kynu-1 mutants……………………………………………...... 39

Figure 3-4: Loss of kynu-1 blocks deuterium label supplied in Trp from being incorporated into

NAD+………………………………………………………………………...…………………. 39

Figure 3-5: Loss of salvage NAD+ synthesis increases of tdo-2 transcript levels……………… 40 ix

Figure 3-6: QA, Kyn and 3HAA supplementation rescues gonad delay in loss of salvage NAD+

synthesis mutants……………………………………………………………………...... 41

Figure 3-7:QA supplementation restores NAD+ levels in loss of salvage NAD+ synthesis

mutants………………………………………………………………………………….. 41

Figure 3-8: Supplementation with QA reverses glycolytic blockage in loss of salvage NAD+

synthesis mutants………………………………………………………………….……. 42

Figure 3-9: umps-1 blocks QA ability to rescue gonad delay…………………………………... 43

Figure 3-10: Loss of umps-1 decreases global NAD+ levels…………………………………… 44

Figure 3-11: Trp pool is isotopically labeled in N2 and umps-1 mutants and isotope label from

Trp is incorporated into QA in both N2 and umps-1 mutants……………...... 45

Figure 3-14: Loss of umps-1 blocks label in Trp from being incorporated into NAD+...... 46

Figure 3-14: Loss of umps-1 blocks QA incorporation into NAD+…………………………….. 47

Figure 3-16: Loss of NAD+ de novo synthesis disrupts fecundity……………………………... 47

Figure 4-1: NMRK-mediated synthesis for NAD+ biosynthetic capacity…………………….. 55

Figure 4-2: Both NR and NA supplementation restore NAD+ in pnc-1 mutants………………. 56

Figure 4-3: Both NR and NA supplementation reverses pnc-1-mediated changes in levels

of glycolytic intermediates……………………………………………………………... 57

Figure 4-4: Loss of nmrk-1 decreases global NAD+ levels……………………………………... 58

Figure 4-5: Loss of salvage NAD+ synthesis results in up-regulated nmrk-1 mRNA

level………………………………………………………………………………….….. 59

Figure 4-6: Embryogenesis is extended for 240 hours in nmrk-1 mutants……………………... 61

Figure 4-7: Embryogenesis is extended for up to 28 hours in clk-1 mutants……………….….. 61

Figure 4-8: Embryogenesis is extended for up to 72 hours in isp-1 mutants…………....……... 62 x

Figure 4-9: Health and life span in nmrk-1 mutants are comparable to N2…………………….. 63

Figure 5-1: NAD/NADH ratio in pnc-1 mutants………………………………………...... 72

Figure 5-2: PARP deletion increases global NAD+ levels in pnc-1 mutants…………...... 73

Figure 5-3: Loss of NMRK-mediated synthesis results in up-regulated tdo-2 and pnc-1 mRNA

levels……………………………………………………………………………………. 74

Figure 5-4: Loss of de novo NAD+ synthesis results in up-regulated pnc-1 and nmrk-1 mRNA

levels……………………………………………………………………………………. 75

Figure 5-5: Loss of kynu-1 results in TCA cycle perturbations………………………………… 76

Figure 5-6: Loss of kynu-1 results in increased glycolysis……………………………………... 77

Figure 5-7: Loss of nmrk-1 results in increased citrate/isocitrate……………………………… 79

Figure 5-8: Loss of nmrk-1 results in increased glycolysis……………………………………. 80

Figure 5-9: Schematic summarizing glycolytic and TCA metabolic changes observed in

loss of de novo and NMRK-mediated synthesis mutants………………………………. 81

Figure 5-10: Loss of NAD+ biosynthetic pathway does not affect oxygen consumption……….82

Figure 5-11: Loss of NAD+ biosynthetic pathway does not affect heat production…………… 83

Figure A-1: Alanine steady state levels are increased in pnc-1 mutants……………………… 111

Figure A-2: α-ketoglutarate and glutamate steady state levels are increased in pnc-1

mutants………………………………………………………………………………… 111

Figure A-3: Alanine aminotransferase mRNA levels are up-regulated in pnc-1

mutants………………………………………………………………………………... 112

Figure A-4: Pyruvate steady state levels in worms treated with agxt-1 RNAi………………... 112

Figure A-5: Metabolic carbon tracing with stable isotope alanine in N2 and pnc-1

mutants………………………………………………………………………………… 113 xi

LIST OF TABLES

Table 2-1: Average % glucose label incorporation in N2 and pnc-1 mutants…………………. 20

Table 2-2: Average % glucose label incorporation in N2 and pnc-1 mutants………………….. 24

xii

LIST OF ABBREVIATIONS

1,3-BPG 1,3-bisphosphoglycerate 3HAA 3-hydroxy anthranilate 3HK 3-hydroxy L-kynurenine 3PGA Glycerate 3-phosphate AGXT-1 Alanine-Glyoxylate aminotransferase AFMD-1 Arylformamidase ATP Adenosine triphosphate CLK-1 Clock (biological timing) abnormality CSB Cockayne syndrome group B DHAP Dihydroxyacetone phosphate FGT-1 Facilitate glucose transporter G3P Glyceraldehyde 3-phosphate GAPDH Glyceraldehyde 3-phosphate dehydrogenase G6P Glucose-6-phosphate HAAO-1 Hydroxyanthranilate 3,4-dioxygenase HDL High-density lipoproteins ISP-1 Iron-Sulfur Protein KMO-1 Kynurenine 3-monooxygenase KYN Kynurenine KYNU-1 kynureninase LigAs NAD+-dependent DNA ligases LDL Low-density lipoprotein NADH Nicotinamide adenine dinucleotide reduced Nampt Nicotinamide phosphoribosyltransferase NA Nicotinic acid NaAD Nicotinic acid adenine dinucleotide NAD+ Nicotinamide adenine dinucleotide NAM Nicotinamide NaMN Nicotinic acid mononucleotide xiii

NaR Nicotinic acid riboside NGMA Nongrowing but metabolically active NFK N-formylkynurenine NMDA N-methyl-D-aspartic acid NMN Nicotinamide mononucleotide NMRK-1 Nicotinamide Riboside Kinase 1 NR Nicotinamide riboside PARP Poly ADP-ribose polymerase PARP-1 Poly ADP-ribose polymerase related PEP Phosphoenolpyruvate PNC-1 Pyrazinamidase and nicotinamidase QA Quinolinic acid QPRTase Quinolinic acid phosphoribosyltransferase TRP Tryptophan TDO-2 Tryptophan/indoleamine 2,3-dioxygenase UMPS-1 Uridine monophosphate synthetase XPA Xeroderma pigmentosum group A

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ACKNOWNLEDGMENTS

All the honor and all the glory belongs to The Most High for choosing me take this ordained journey. I can never repay You, God, for what You done for me. How You loosed my shackles and You set me free. How You made a way out of no way. Turned my darkness into day. You’ve been my joy in the time of sorrow. Hope for my tomorrow. Peace in the time of storm. Strength when I’m weak and worn. All for Your will and Your glory, I am forever grateful and thankful that I was chosen and the destiny that’s been bestowed upon me.

I am forever giving gratitude to all the ancestors who overcame tremendous obstacles, but stayed resilient, so that I could have this opportunity today. For that, I will touch the sky. My grandmother, Isabella Yarbrough Miller, and great-aunt, Minnie Pearl Yarbrough, from the very beginning believed in me and told me I could conquer whatever my imagination could dream.

When I was the age of five, these women asked, what would I be when I grew up and what would I be remembered for? I responded I would be a famous scientist. But God. My mother has been my biggest role model. I grew up witnessing this woman touch and change lives on a daily basis through education, activism and empowerment. My wish is that I can have just half of an impactful career touching and saving lives like my mother. Countless people have supported me, while overlooked by many more; to the ones who believed me in and to the ones who did not, watch while I change the world.

The last seven years have truly been an adventure and journey. I still rejoice and thank

God for every step of the process. I am forever grateful to the wonderful mentors that were strategically placed in my path to train me, and give me the guidance needed to grow more.

Being able to train under Wendy was truly God-ordained. The world of science is there for me to conquer, because Wendy saw something great in me and had the patience to mold me into the xv scientist I’ve become today. The atmosphere at Penn State allowed me to fail and make mistakes, but also allowed me to be great. I was able to grow and flourish not only as a scientist but also as a leader. It’s the little things that counts and really makes a world of difference.

Lastly, I’m forever greatly for the ones I went through this journey with. There is nothing like getting your PhD with your best friends. We started out as strangers, but finished as family.

Together we made it, but we already know that we are better together. Let’s continue on our

God-ordained purpose to influence and change this world for the glory and will of God.

1

Chapter 1

Introduction

Part I: NAD+ is a central hub in cellular metabolism

Over the last century, our society has gone from experiencing diseases associated with

NAD+ deficiency to targeting NAD+ biosynthesis and metabolism for therapeutic potential.

Although the dietary requirement for vitamin B3 is well known, the impact of manipulating

NAD+ biosynthesis and metabolism remains understudied. This dissertation focuses on the biochemical and biological impact of NAD+ biosynthesis. Thus, I will review aspects of the role of NAD+, its discovery and future use.

+ Historical context of Vitamin B3 as a precursor for NAD

A central metabolic cofactor in eukaryotic cells, nicotinamide adenine dinucleotide

(NAD+), plays a vital and critical role in energy metabolism regulating cellular metabolism and energy homeostasis. NAD+ was first described as a cofactor involved in fermentation (Harden and Young, 1906). In 1906, researchers Harden and Young discovered cozymase from yeast extract. They observed boiled and filtered yeast extract facilitated alcoholic fermentation in yeast extract that was not boiled. Cozymase was the mixture of components responsible for this reaction (Harden and Young, 1906). In 1923, cozymase was purified and identified as a nucleoside sugar phosphate, essential for carbohydrate usage (Euler-Chelpin, Nobel Lecture,

1929). Finally, NAD+ was isolated from cozymase by Otto Warburg, and its role in hydrogen transfer in biochemical reactions was discovered (Koppenol, Bounds, & Dang, 2011).

+ Vitamin B3 is the active precursor for NAD biosynthesis. Vitamin B3 deficiency is often associated with diets that are low in protein (Bogan & Brenner, 2008a; Chi & Sauve, 2013; A. A.

Sauve, 2008). This was first identified two centuries ago in farmers whose diet depended on 2

maize and was poor in meat (Rajakumar, 2000). Chronic lack of dietary Vitamin B3 and tryptophan, both precursors of NAD+, serves as the cause of pellagra (Bogan & Brenner, 2008a;

Rajakumar, 2000; A. A. Sauve, 2008). This disease is associated with dermatitis, diarrhea and dementia. Recently, it was as an epidemic in the southern United States during the 1900s, and its common in malnourished communities of rural Europe (Sydenstricker, 1958.). Interestingly, niacin is found in maize; however, it is not bioavailable unless the maize undergoes an alkali treatment. Aztecans and Mesoamericans used this process to treat maize (Gwirtz & Garcia-Casal,

2014.). NAD+ is not only important for the prevention of pellagra, but is also associated with extended lifespan, increased resistance against infectious and inflammatory diseases (Canto,

Menzies, & Auwerx, 2015; Sauve, 2008) and is likely very important in resisting a number of other disease processes (Xu & Sauve, 2010) such as cardiovascular disease, metabolic syndrome, neurodegenerative diseases and even cancer.

Role of NAD+ in redox reactions

NAD+ and its reduced and phosphorylated forms, functioning as an oxidoreductase cofactor in a wide range of metabolic reactions, are vital in cellular metabolism regulation and energy production. In its reduced form, NADH serves as the primary electron donor in the mitochondrial respiratory chain producing ATP, the currency of energy transfer. NAD+ pools can modulate compartment-specific metabolic pathways activity. In the cytosol this includes glycolysis, and in the mitochondria this includes the TCA cycle, oxidative phosphorylation, fatty acid and amino acid oxidation (Cerutti et al., 2014; French et al., 2010).

Glycolysis requires two NAD+ molecules to convert glucose into pyruvate. The production of lactate from pyruvate oxidizes NADH back into NAD+ in the cytosol. NADH equivalents generated via glycolysis in the cytosol are transferred to the mitochondria by the 3 malate-aspartate shuttle. The TCA cycle reduces NAD+ molecules to produce multiple NADH molecules. NADH from glycolysis or the TCA cycle can react at Complex I in the electron transport chain (Sazanov, 2015). Because cellular NAD+ pools can be limiting, both glycolysis in the cytosol and the TCA cycle in the mitochondria can influence metabolic homeostasis by altering NAD+ and NADH levels (Bai et al., 2011; Pirinen et al., 2014; Pittelli et al., n.d.). This illustrates the importance of compartmental and tissue specific NAD+ requirements to maintain cellular and metabolic homeostasis.

Role of NAD+ as a substrate

NAD+ is not only a vital cofactor/coenzyme involved in key redox reactions, but it also serves as a signaling messenger that can modulate metabolic and transcriptional responses in cells. NAD+ as a substrate has been implicated in a wide array of biological functions. NAD+ consumption is linked to activities that include control of cellular metabolism and energy homeostasis, aging and longevity, transcriptional silencing, cell survival, proliferation, differentiation, DNA damage response, stress resistance and apoptosis (Hasmann & Schemainda,

2003; Hegyi, Schwartz, & Hegyi, 2004; Rowent, 1951; Williams, Jones, & Agarose, 1985; T.

Yang, Chan, & Sauve, 2007). Sirtuins, PARPs and CD38 serve as the key enzymes that consume

NAD+ for their reactions.

Sirtuins serve as metabolic sensors of cells as their activity is coupled to changes in the cellular NAD+/NADH redox state (Canto et al., 2015). They are activated in response to nutrient deprivation and energy deficit (Bonkowski & Sinclair, 2016; Haigis, Sinclair, & Edu, n.d.). This triggers cellular adaptations, which leads to improved metabolic efficiency in stressed conditions

(Bonkowski & Sinclair, 2016; Haigis et al., n.d.). PARP activity constitutes the main NAD+ catabolic activity (Bai et al., 2011; Chiarugi, 2012; Mouchiroud, Houtkooper, & Auwerx, 2013; 4

Y. Yang & Sauve, 2016). PARPs are activated in response to DNA damage and genotoxic stress

(Bai et al., 2011; Chiarugi, 2012). However, the CD38 family of bi-functional enzymes uses

NAD+ to generate cADP-ribose, which serves as an intracellular second messenger for calcium signaling (Guse, 2005). NAD+ metabolism triggering of major signaling processes has sparked interest in the dynamics that regulate these mechanisms.

Part II: Eukaryotic NAD+ biosynthesis

Overview of NAD+ biosynthesis

A critical balance between NAD+ biosynthetic and consuming pathways establishes the

NAD+ cellular pool. As NAD+ consumer enzymes consume NAD+ for their reactions, organisms must have the means to replenish the NAD+ pool via de novo or salvage synthesis (Rongvaux,

Andris, Van Gool, & Leo, 2003). There are four major molecules that serve as precursors for

NAD+ biosynthesis, and these compounds can be taken up from the diet. The half-life for intracellular NAD+ is short, and NAD+ is not distributed evenly in subcellular compartments

(Ying, 2008; Guillemin, et., 2003). Therefore, there’s a critical need to maintain global NAD+ homeostasis. Furthermore, this suggests that there are also unique mechanisms in place to facilitate biosynthetic capacity requirements.

Salvage NAD+ synthesis: In a majority of species, salvage synthesis is the main source of

NAD+ from dietary niacin (French et al., 2010; Preiss and Handler, 1957; Srivastava, 2016).

Salvage biosynthesis of NAD+ is required to replenish NAD+ by the conversion of nicotinamide

(NAM) into nicotinic acid (NA), where it enters back into the Preiss-Handler pathway (Tracy L

Vrablik, Wang, Upadhyay, & Hanna-Rose, 2011). It’s imperative for salvage synthesis to occur because NAM is a noncompetitive inhibitor of NAD+ consumers (Belenky, Bogan, & Brenner,

2007). In mammals, Nampt converts NAM into NAD+ (H. Yang, Lavu, & Sinclair, 2006), and in 5

C. elegans, the nicotinamidase, PNC-1, converts NAM into NA (T L Vrablik, Huang, Lange, &

Hanna-Rose, 2009). Although Nampt and nicotinamidases are enzymatically different, they both promote NAD+ synthesis by consuming NAM, suggesting functional equivalence (H. Yang et al., 2006). Consistent with this evidence, we previously reported that Nampt could partially substitute for PNC-1 in C. elegans (T L Vrablik et al., 2009).

Preiss-Handler Pathway: The highly conserved Preiss-Handler pathway was the first pathway to be intensively studied, which produces NAD+ from either NA. NA or QA is converted to nicotinic acid mononucleotide (NaMN) by transfer of a phosphoribose moiety.

Next, an adenylate moiety is transferred to form nicotinic acid adenine dinucleotide (NaAD).

Finally, the nicotinic acid moiety in NaAD is amidated to a nicotinamide (NAM) moiety, forming nicotinamide adenine dinucleotide (Preiss and Handler, 1957). NA was first used for its ability to lower cholesterol levels and treatment against dyslipidemia (Altschul, Hoffer, &

Stephen, 1955). NA reduced plasma triglyceride and LDL levels, while increasing HDH levels.

However, the use of NA in the clinic has been limited due to continuous flushing (Birjmohun,

Hutten, Kastelein, & Stroes, 2005). Activation of a G protein couple receptor was the reason of flushing in patients, rather than driving NAD+ synthesis (Benyo et al., 2005.).

de novo NAD+ synthesis: NAD+ can be synthesized via the de novo synthesis pathway from the amino acids aspartate, in prokaryotes, and tryptophan, in eukaryotes (A. a Sauve, 2008).

Tryptophan is converted into NAD+ through an eight-step pathway, oxidizing Trp through the kyneurine pathway and finally producing QA, which enters the Preiss-Handler pathway (Bogan

& Brenner, 2008a). The nutritional requirement for ongoing de novo synthesis is necessary for the prevention of pellagra and the progression of NAD+ biosynthesis (Belenky et al., 2007).

Tryptophan is one of the essential amino acids that the human body is incapable of 6 synthesizing. The kynurenine pathway is the major route for tryptophan metabolism. As tryptophan proceeds through the kynurenine pathway to achieve the final product, NAD+, several neuroactive intermediates are generated. These intermediates consist of the free-radical generator, 3-hydroxyanthranilic acid (3HAA) (Goldstein et al., 2000), the excitotoxin and N- methyl-D-aspartic acid (NMDA) receptor agonist, quinolinic acid (QA) (Stone & Perkins, 1981), the NMDA antagonist, kynurenic acid (Perkins & Stone, 1982) and the neuroprotectant, picolinic acid (Jhamandas, Boegman, Beninger, & Bialik, 1990). An accelerated degradation of tryptophan with an accompanied increase in kynurenine metabolites in the serum, CSF and brain tissue is often associated with various pathological conditions (Chen & Guillemin, 2009; Sas,

Robotka, Toldi, & Vécsei, 2007a). Up-regulation of the kynurenine pathway is associated with infectious diseases, neurological disorders, affective disorders, autoimmune diseases, peripheral conditions and malignancy (Erhardt, Schwieler, Imbeault, & Engberg, 2016; Fatokun, Hunt, &

Ball, 2013; Lim et al., 2015; Majewski, Kozlowska, Thoene, Lepiarczyk, & Grzegorzewski,

2016; O’Farrell & Harkin, 2017; Ohashi, Kawai, & Murata, 2013; Oxenkrug, 2010; Vamos,

Pardutz, Klivenyi, Toldi, & Vecsei, 2009). Therefore, it is imperative to elucidate the relationship between NAD+ synthesis and the kynurenine pathway. The C. elegans’ genome encodes all the enzymes involved in the kynurenine pathway; however, the genome lacks the critical quinolinic acid phosphoribosyltransferase (QPRTase) homolog that converts QA into

NaMN for biosynthesis of NAD+. Therefore, the presence of this pathway in C. elegans has been in questioned.

Riboside synthesis: Nicotinamide riboside (NR) was recently discovered as an additional

+ vitamin B3 precursor for NAD biosynthesis (Bieganowski & Brenner, 2004). This vitamin is naturally found in cow’s milk (Bieganowski & Brenner, 2004). NR is converted into NAD+ via 7 the NRK pathway (Belenky et al., 2007). This pathway can also be used for nicotinic acid riboside (NaR) salvage in yeast and mammalian cells (Bogan & Brenner, 2008b; Ratajczak et al.,

2016; Trammell et al., 2016). NR has been shown to be a NAD+ precursor in yeast and mammalian cells, by contributing to maintenance of NAD+ levels (Belenky et al., 2007;

Bieganowski & Brenner, 2004; Ratajczak et al., 2016; Trammell et al., 2016). Also, NR has been shown to improve longevity and gene silencing in yeast (Bogan & Brenner, 2008a), along with an array of conditions in mammalian models, such as neurological disorders, metabolic syndromes, cancer and aging (Mills et al., 2016). It’s been suggested that the only vitamin B3 precursor supporting neuronal NAD+ synthesis is NR (Belenky et al., 2007; Bogan & Brenner,

2008a). Interestingly, NRK is highly conserved from yeast to humans (Belenky et al., 2007).

NAD+#Biosynthesis#in#C.'elegans'' Trp#

Kynurenine# de'novo' pathway## synthesis#

QA# ???#

NaMN#

Salvage# NaAD# Synthesis# NA#

NRK# pnc%1'

Pathway# qns%1' nmrk%1' NR# NMN# NAD+# NAM# NAD+#Consumers# Sirtuins# PARPs#

Figure 2-1: NAD+ biosynthesis in C. elegans Schematic outlining NAD+ biosynthetic pathways in the nematode, C. elegans. Salvage NAD+ synthesis is represented in blue, NMRK-mediated synthesis is represented in orange and de novo NAD+ synthesis is represented in green. NAD+ consumers, sirtuins and PARPs are outlined in gray.

8

Part III: Targets for NAD+ biosynthesis and metabolism based drug discovery

Over the last couple of decades, NAD+ has emerged as a critical link bridging regulatory and bioenergetics processes (Srivastava, 2016). NAD+ functions in a majority of metabolic pathways, suggesting NAD+ limitations would perturb metabolic efficiency. NAD+ has the ability to respond dynamically to physiological stimuli. It’s proposed that decreased NAD+ levels promote development of ailments associated with aging and disease (Mills et al., 2016). Hence,

NAD+ biosynthesis and metabolism has proven to be an attractive and promising therapeutic target for influencing health-span and obesity-related phenotypes as well as tumor growth.

However, the impact of manipulating NAD+ biosynthesis and metabolism on metabolic homeostasis remains understudied.

Cancer inhibition

NAD+ is a key metabolite that’s required for tumor growth and progression (Srivastava,

2016). Elevated NAD+ levels, both through PARP inhibition or NAD+ precursor supplementation, can rewire cellular metabolism and enhance oxidative versus glycolytic metabolism (Bai et al., 2011; Chiarugi, 2012). Remodeling metabolism, by enhancing NAD+ levels, could potentially constitute a complementary mechanism to slow down cancer progression or initiate cell death (Canto et al., 2015; Srivastava, 2016). However, depletion of

NAD+ is proposed to inhibit growth of several cancer models (Canto et al., 2015; Srivastava,

2016). A majority of cancer cells rely on glycolytic metabolism versus oxidative phosphorylation. Because a majority of cancer cells rely on increased central carbon metabolism and biomass production to sustain unrestricted growth, reducing NAD+ bioavailability is reported to have antineoplastic effects in tumor cells (Chiarugi et al., 2012; Tateishi et al., 2015). In addition to the negative impact on metabolic rearrangements that fuel cancer growth, decreasing 9

NAD+ can also halt tumor progression by limiting NAD+-dependent enzymes activity that promotes tumor growth (Houtkooper & Auwerx, 2012; Srivastava, 2016). Therefore, it’s conceivable that limiting NAD+ availability would counteract the metabolic changes that promote cancer growth and progression. This suggests that both boosting and depleting NAD+ biosynthesis can impact tumor development and progression.

At the root of all cancers is genomic stress. Therefore, an essential tool for preventing cancer is maintaining genome integrity. NAD+ consumers, PARPs and sirtuins, both have key roles in maintaining genomic integrity (Bai et al., 2011; Chiarugi, 2012; Haigis et al., n.d.; Y.

Yang & Sauve, 2016). This suggests NAD+ regulation could impact cancer susceptibility and development. PARP inhibitors are currently in phase III clinical trials as anti-cancer agents, because these compounds enhance oxidative metabolism and improve metabolic flexibility (Y.

Yang & Sauve, 2016). In contrast, the ability to target sirtuins for anti-cancer benefits is still being worked out. SIRT1 protects against age-related carcinomas and sarcomas in transgenic animal models, but not lymphomas (Herranz et al., 2010).

Niacin deficiency enhances susceptibility to cancer development and progression; this suggests that cellular NAD+ levels are inversely related to the incidence of cancer (Benavente,

Schnell, & Jacobson, 2012). Supplementation with niacin can decrease the development of skin cancer (Gensler, Williams, Huang, & Jacobson, 1999). Also, increasing NA and NAM precursors can inhibit metastasis and breast cancer progression (Santidrian et al., 2013).

Interestingly, NR can both reduce the incidence of cancer and have a therapeutic effect on fully formed tumors in a mouse model with liver cancer (Tummala et al., 2014). Furthermore,

NAMPT is overexpressed in several tumor models and is associated with tumor progression (Bi et al., 2011; Hasmann & Schemainda, 2003; Van Beijnum et al., 2002; Wang et al., 2011). NAD+ 10 depletion by down-regulating NAMPT activity can reduce tumor cell growth and sensitize cells to chemotoxic agents (Bi et al., 2011; Hasmann & Schemainda, 2003; Wang et al., 2011; Watson et al., 2009).

Health and lifespan benefits

With the increased trend of the population of people living longer, there has been an increase in promoting healthier aging to reduce costs associated with aging and disease (Mills et al., 2016). Mitochondrial dysfunction is a hallmark for aging and diseases, such as diabetes, obesity, neurodegeneration and cancer (Srivastava, 2016). Reduced NAD+ levels are strongly implicated in these disorders (Cerutti et al., 2014; Houtkooper & Auwerx, 2012; Khan et al.,

2014; Mouchiroud et al., 2013). Reductions in NAD+ have been consistently observed during aging in worms, diverse rodent tissues and human skin samples (Braidy et al., 2011; Gomes et al., 2013; Khan et al., 2014; Massudi et al., 2012; Mouchiroud et al., 2013; Yoshino, Mills,

Yoon, & Imai, 2011). The declining NAD+ pool during aging ultimately compromises mitochondria function in these models, which can be restored with NAD+ precursor supplementation or inhibition of PARP activity (Gomes et al., 2013; Mouchiroud et al., 2013;

Zhang et al., 2016). Also, metabolic disturbances in mice caused by high fat diets are corrected with boosting NAD+ levels (Baur et al., 2006; Canto & Auwerx, 2012; Lagouge et al., 2006;

Yoshino et al., 2011). Mice engineered to express additional copies of SIRT1 or SIRT 6, or treated with sirtuin-activating compounds or NAD+ precursors, have improved organ function, physical endurance, disease resistance and longevity (Bonkowski & Sinclair, 2016). Proper function of the mitochondria is critical for the maintenance of metabolic homeostasis and activation of appropriate stress responses.

11

Neurological disorders

Both NAD+ precursors and PARP inhibitors are known to be neuroprotective.

Interestingly, there is an induction of multiple transcripts for NAD+ biosynthetic enzymes following injury of neurons. In this response, there is more than a 20-fold increase in NRK2 mRNA levels (Sasaki, Araki, & Milbrandt, 2006), which catalyzes the synthesis of NAD+ from

NR (Bieganowski & Brenner, 2004; Chi & Sauve, 2013; Ratajczak et al., 2016). This suggests a compensatory response to raise NAD+ levels during neural injury. Further supporting this observation, pretreatment of neurons with NAD+, NR and NMN can protect against axonal degeneration and hearing loss in mice (Brown et al., 2014; Gerdts, Brace, Sasaki, Diantonio, &

Milbrandt, 2015; Sasaki et al., 2006; L. Wang, Ding, Salvi, & Roth, 2014). Modeling protein misfolding in Alzheimer’s and Parkinson’s disease, by exposing neuronal cells to toxic prion proteins, induced NAD+ depletion that was improved by exogenous NAD+ and NAM (Zhou et al., 2015). Moreover, increasing NAD+ biosynthesis via NAM pharmacological doses provides protection against ischemia (Klaidman et al., 2003; Sadanaga-Akiyoshi et al., 2003), fetal alcohol-induced neurodegeneration (Ieraci & Herrera, 2006) and fetal ischemic brain injuries (Y.

Feng, Paul, & LeBlanc, 2006) by preventing NAD+ depletion in rodent models. Supplementation with NR improves Alzheimer’s disease phenotype via PGC-1α-mediated β-secreatase (BACE1) degradation and induction of mitochondrial biogenesis (Gong et al., 2013), illustrating that increased NAD+ levels can attenuate increase in β-amyloid content and oxidative damage. This increased in NAD+ levels prevented cognitive decline and neurodegeneration in rodent models of

Alzheimer’s disease (Gong et al., 2013; Qin et al., 2006; Turunc Bayrakdar, Uyanikgil, Kanit,

Koylu, & Yalcin, 2014). The depletion of NAD+ in neurodegeneration is generally attributed to the activation of PARP enzymes. In DNA repair disorders, such as xeroderma pigmentosum 12 group A (XPA) and Cockayne syndrome group B (CSB), PARP-1 activation occurs. Treatment with specific PARP inhibitors can reverse the defective phenotypes in XPA mutant worms and

CSB mutant mice respectively (Fang et al., 2014; Scheibye-Knudsen, Fang, Croteau, & Bohr,

2014). Therefore, it’s proposed that maintaining NAD+ levels and homeostasis sustains basal metabolic function and health in neurons.

Novel antibiotics

Due to the alarmingly and growing reports of bacterial resistant strains to antibiotics, there’s a critical and urgent need for new targets and potential antibacterial agents (Kim et al.,

2013; Murima, McKinney, & Pethe, 2014; Pankiewicz, Petrelli, Singh, & Felczak, 2015). Over the last 30 years, there has been a worldwide problem with multiple drug resistance to antibiotics among pathogenic bacteria spreading (Murima et al., 2014). Therefore, there’s been an increased push to search for novel inhibitors with distinct modes of action from diverse chemical classes

(Pankiewicz et al., 2015). DNA ligases are essential for cellular and biochemical processes, including DNA replication, recombination and repair (Gu et al., 2012; Stokes et al., 2012). These enzymes have been identified as an attractive antibacterial drug target (Harris & Pierpoint, 2012;

Timson, Singleton, & Wigley, 2000). Based on substrate requirements, DNA ligases are divided into two classes, NAD+ or ATP dependent. NAD+-dependent ligases are highly conserved in bacteria, whereas, ATP-dependent ligases are present in eukaryotic cells. NAD+-dependent DNA ligases (LigAs) are essential enzymes in bacteria and vital for DNA replication and repair

(Shuman, 2009). Nongrowing but metabolically active (NGMA) bacteria are known to be a refractory subpopulation to current antibacterial (Murima et al., 2014; Rittershaus, Baek, &

Sassetti, 2013). Targeting NAD+ biosynthesis eliminates NGMA cells and impairs the establishment of bacterial infection (Kim et al., 2013). Therefore, because LigA is essential for 13 bacteria, and is absent in eukaryotic cells, it is an attractive therapeutic target for the development of broad-spectrum antimicrobial agents.

An#$Cancer*

Novel* NAD+* Health*and* An#bio#cs** lifespan*

Neurological*Disorders*

Figure 1-3: Therapeutic targets for NAD+ biosynthesis and metabolism Schematic representing that both boosting and inhibiting NAD+ biosynthesis and metabolism can be therapeutic based targets for diseases.

Part IV: Role of NAD+ biosynthesis in metabolic homeostasis

The broad spectra of responsibilities for NAD+ in energy production and cellular metabolism, indicates the importance maintaining NAD+ homeostasis. Tissue and compartmental-specific NAD+ depletion has been associated with various disease and aging aliments. Therefore, NAD+ biosynthetic capacity is vital to maintain metabolic homeostasis in organisms. The biochemistry of NAD+ metabolism and NAD+ biosynthetic pathways are known; however, the biological impacts upon inhibition or boosting of these pathways are not known.

The mechanisms that alter NAD+ metabolism include multiple processes, but the scope of these mechanisms is currently very unclear. This work seeks to elucidate the developmental and physiological roles of NAD+ biosynthetic pathways in C. elegans. 14

Due to the requirement and capacity of NAD+ pools in cellular metabolism, I hypothesize that there are homeostatic interactions amongst these pathways. Also, I propose that there are unique and diverse biological functions for each NAD+ biosynthetic pathway. Within the last

+ couple of decades, NR was discovered to be a newly found Vitamin B3 precursor for NAD biosynthesis. This suggests that there could still be unknown routes and mechanisms for NAD+ biosynthesis. NAD+ biosynthesis and metabolism is an attractive therapeutic target for a growing number of disease states. However, it’s imperative to elucidate the biological impact, homeostatic interactions and metabolic perturbations of manipulating NAD+ biosynthesis. C. elegans are a great model to define the developmental and physiological roles of NAD+ biosynthesis in a multicellular organism. This work uncovers unique biological functions for

NAD+ biosynthetic pathways, a compensatory network amongst pathways and underground metabolism in NAD+ biosynthesis. This supports the dynamic role of NAD+ biosynthesis and metabolism in cellular homeostasis.

15

Chapter 2

Metabolic carbon tracing reveals disrupted glycolysis due to a loss of salvage NAD+ synthesis in C. elegans

Introduction

Importance of NAD+ in energy-producing redox reactions

As a key link connecting regulatory and bioenergetics process (Chiarugi, 2012), NAD+ has emerged as a central metabolic co-factor playing a critical role regulating cellular metabolism and energy homeostasis (Srivastava, 2016; Y. Yang & Sauve, 2016). NAD+ provides a vital role within energy metabolism, accepting hydride equivalents to form reduced NADH.

This reaction furnishes reducing equivalents to the mitochondria to fuel oxidative phosphorylation via the electron transport chain. Due to the critical nature of this co-enzyme in energy-producing pathways, NAD+ serves as a hub driving cellular metabolism.

The oxidized and reduced form of NAD+ is crucial for the function of numerous metabolic redox reactions in all species. The NAD/NADH ratio is responsible for regulating the activity of metabolic pathways such as glycolysis, citric acid cycle and fatty acid oxidation. The requirement of NAD+ in these reactions suggests that NAD+ limitations would likely perturb metabolic efficiency and homeostasis. NAD+ pools in either the cytosol or mitochondria can modulate the activity of compartment-specific metabolic pathways. For example, NAD+ pools in the cytosol modulate glycolysis, whereas the TCA cycle and oxidative phosphorylation are modulated by mitochondria’ NAD+ pools. The importance of NAD+ in central metabolism suggests that disruptions to NAD+ levels will contribute to the development of ailments associated with aging, health and disease.

16

Relationship between glucose metabolism and cellular NAD+ pools

Otto Warburg was the first to isolate NAD+ and show its role in hydrogen transfer

(Canto, Menzies, & Auwerx, 2015; Chiarugi et al., 2012; Koppenol, Bounds, & Dang, 2011;

Yang & Sauve, 2016). Based on his studies, Warburg suggested that lactate production was increased in tumor cells, indicating an accelerated rate of glycolysis in order to facilitate both biomass production and oxidative phosphorylation. Therefore in tumor cells, the cytosolic NAD+ population is being regenerated from NADH via reducing pyruvate to lactate, instead of eventually transferring electrons from NADH to the mitochondrial respiratory chain.

Two molecules of NAD+ per molecule of glucose are required for conversion of glucose into pyruvate. Glyceraldehyde 3-phosphate dehydrogenase is the key enzyme using NAD+ in the sixth step of glycolysis where glyceraldehyde 3-phosphate (G3P) is oxidized to 1,3- bisphosphoglycerate (1,3-BPG) in the cytosol. The NADH equivalents generated from this reaction in the cytosol are transferred to the mitochondria via the malate-aspartate shuttle.

Because cellular NAD+ can be limiting, altering cytosolic and nuclear NAD+ and NADH levels can influence metabolic homeostasis across all cellular compartments (Bai et al., 2011; Pirinen et al., 2014; Pittelli et al., n.d.). Regulating NAD+ metabolism and biosynthesis seems to be quite promising for promoting healthier aging and treating several disease models (Mills et al., 2016).

However, the impact of manipulating NAD+ biosynthetic pathways remains understudied.

Understanding the alterations that occur to the developmental and physiological system amongst perturbations to NAD+ homeostasis is critical to move closer to uncovering the therapeutic benefits of this metabolite.

17

Loss of NAD+ salvage synthesis disrupts glycolysis leading to developmental reproductive delay in C. elegans

Most organisms rely on NAD+ salvage synthesis as their main source of NAD+ biosynthetic capacity (Srivastava, 2016). Therefore, successfully recycling NAM, released from

NAD+ consuming reactions, back to NAD+ is a major challenge for maintaining NAD+ metabolism and homeostasis. The first step in C. elegans’ salvage NAD+ synthesis involves the nicotinamidase, pnc-1, which is responsible for the conversion of NAM into NA (T L Vrablik et al., 2009). NA enters the Preiss-Handler pathway where it ultimately is processed into NAD+.

We previously linked loss of salvage NAD+ synthesis to disruptions in glycolysis, which lead to reproductive development delay (W. Wang et al., 2015). Global metabolic profiling suggested glycolytic blockage in our pnc-1 mutants. Steady state levels of metabolites above the step using

NAD+ to convert G3P into 1,3BPG were increased, and the steady state levels of metabolites after this step were decreased in pnc-1 mutants (Figure 2-1). To investigate if these metabolic changes were functionally relevant to the developmental gonad delay phenotype, we supplemented pnc-1 mutants with 3PGA and PEP and were able to rescue the gonad delay phenotype observed in loss of salvage synthesis mutants. Although glycolysis was apparently blocked in our pnc-1 mutants, pyruvate levels were inconsistent with the glycolytic trend. Also, the citric acid cycle was not perturbed and functions of the mitochondria were intact in pnc-1 mutants (W. Wang et al., 2015). This observation suggested that loss of salvage NAD+ biosynthesis is cytosolicic specific. Based on these results, we hypothesized that glycolysis is blocked at the NAD+-dependent step due to loss of salvage NAD+ synthesis. To gain a deeper understanding of the core mechanisms behind NAD+ salvage synthesis and glucose metabolism, we decided to directly test this model via application of metabolic isotopic carbon tracing tools.

Metabolic carbon tracing confirmed that glycolysis is blocked in pnc-1 mutants. Additionally, I 18 observed an increase of isotopic label from pyruvate entering the TCA cycle in these mutants.

Although trehalose steady state levels are increased in pnc-1 mutants, we did not observed an increase of glucose being isotopically shunted towards trehalose. Metabolic carbon tracing provided key insight into elucidating compromised glycolysis due to loss of salvage NAD+ synthesis in C. elegans.

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Figure 2-1: Schematic of disrupted glycolysis in pnc-1(pK9605) mutants. We hypothesized that glycolysis is blocked at the NAD+ dependent step (circled in blue) converting G3P into 1,3BPG in loss of salvage NAD+ synthesis mutants. Steady state levels of metabolites above this step are increased (up arrows), whereas, the levels are decreased in metabolites after this step (down arrows). This specific perturbation was further linked to impaired reproductive development in pnc-1 mutants.

19

Results

Development of metabolic tracing protocol to model compromised glycolysis in pnc-1 mutants

It was both remarkable and intriguing to discover that loss of salvage NAD+ synthesis was associated with impaired reproductive development and compromised glycolysis. To directly test this hypothesis, we decided to use metabolic carbon tracing. First, I optimized a metabolic tracing protocol in C. elegans (Falk et al., 2011; Schrier Vergano et al., 2014). We

13 used universally labeled glucose ( C6-Glucose) as our metabolic tracer in wild-type animals and pnc-1 mutants. Below I have outlined the protocol optimized for metabolic carbon tracing within our research group.

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Figure 2-2: Scheme for metabolic carbon tracing in C. elegans. Approach and protocol optimized to perform metabolic carbon tracing in C. elegans is outlined in detail above. Six independent biological replicates were conducted for both N2 and pnc-1 mutants. 20

Metabolic tracing supports compromised glycolysis in pnc-1 mutants

Loss of salvage NAD+ synthesis results in compromised glycolysis (W. Wang et al.,

2015). Steady state levels of glucose, G6P and DHAP are increased in pnc-1 mutants, and 3PGA and PEP levels are decreased. Therefore, we hypothesized that glycolysis is being blocked at the step using NAD+ to convert G3P to 1,3BPG due to loss of NAD+ salvage synthesis. First, we wanted to investigate the accuracy of our compromised glycolysis model with the optimized metabolic isotopic carbon tracing protocol. We exposed both N2 and pnc-1 mutants to

13 universally labeled glucose ( C6-Glucose) for four hours, and were able to detect isotope label from glucose being incorporated into glycolytic intermediate metabolites, lactate and the TCA cycle metabolites (Tables 2-1 and 2-2). Out of the total glucose pool, a 9% fraction is isotopically labeled after 4 hours of exposure in wild-type animals. However, only 4% of the total glucose pool is isotopically labeled in pnc-1 mutants after 4 hours (Figure 2-4).

Consequently, we next analyzed the flow of isotopic label from glucose through each subsequent step of glycolysis by normalizing each step to the metabolite before, beginning with glucose in both wild-type animals and pnc-1 mutants. Supporting our hypothesis, pnc-1 mutants exhibited an increase of isotope label from glucose in DHAP relative to G6P and a significant decrease in

3PGA relative to DHAP (Figure 2-5). Thus, metabolic carbon tracing supported glycolytic blockage at the expected step, demonstrating glycolysis is compromised due to the loss of salvage NAD+ synthesis.

Avg. % Incorporation Glucose G6P DHAP 3PGA PEP Pyruvate Lactate N2 8.7±7.1 1.3±1.6 5.4±3.6 6.9±4.6 6.6±4.4 5.8±3.9 6.6±4.5 pnc-1 4.1±2.4 0.7±0.5 4.5±2.0 3.7±2.0 4.3±2.1 3.7±1.5 4.1±1.4

Table 2-1: Average % glucose label incorporation in N2 and pnc-1 mutants. Shown are LC-MS measurements of observed % incorporation ± S.D. of isotopically labeled glucose into glycolysis metabolites and lactate in N2 and pnc-1 animals. 21

N2 + isotope Glucose

N2 + isotope +6 shift Glucose +6

N2 control Glucose

N2 control +6 shift Glucose +6 No peak

RT m/z

Figure 2-3: Representative raw data for glucose and isotopic glucose peaks. Glucose and glucose +6 peaks represented with retention time (RT) and mass to charge ratio (m/z) in wild-type worms. This approach was used to view label incorporation into subsequent metabolites.

22

Figure 2-4. Fraction of total glucose pool labeled decreases in pnc-1 mutants. (A). Dot-plot of the % of glucose isotopically labeled in N2 and pnc-1 mutants. (B). Ratio of % glucose incorporated in pnc-1 mutants compared to N2. **, 0.001

Figure 2-5: Glycolysis is blocked at the NAD+ dependent step in pnc-1 mutants. Normalized % isotopic-glucose label incorporated ratios to the metabolite immediately before of pnc-1 mutants compared to N2 animals. p=0.0608, calculated with repeated measures anova. a=n.s, b p=0.05, calculate with anova posttest, Tukey’s multiple comparison test.

Although glycolysis is compromised in pnc-1 mutants, we observed that pyruvate steady state levels, the TCA cycle and mitochondria function were not perturbed in this model (W. 23

Wang et al., 2015). Our metabolic carbon tracing data also revealed that the flow of isotopic label to pyruvate after the NAD+-dependent step is consistent in pnc-1 mutants as wild-type animals (Figure 2-5). We next asked what is the fate of isotopic label from glucose after it reaches pyruvate in loss of salvage NAD+ synthesis mutants. Interestingly, isotopic label in lactate in pnc-1 mutants was comparable to wild-type animals (Figure 2-6). Furthermore, I detected an increase of 50% in isotopic label from glucose entering the TCA cycle in our pnc-1 animals (Figure 2-7). Based on our previous data (W. Wang et al., 2015), we predicted no disruption for the isotopic flow through the TCA cycle in our pnc-1 mutants. Although there is an increase in isotopic flow from pyruvate to citrate, the flow through the subsequent metabolites are similar to wild-type animals in our pnc-1 mutants (Figure 2-7 and Table 2-2), suggesting homeostatic parameters maintaining the cellular pyruvate pool. This supports cytosolic specific loss of salvage NAD+ synthesis disrupts normal glycolytic flux, thus confirming our compromised glycolysis model in pnc-1 mutants.

Figure 2-6: Flow rate of isotopic label after pyruvate into lactate is equal to N2 in pnc-1 mutants. Normalized % isotopic-glucose label incorporated ratios to the metabolite immediately before of pnc-1 mutants compared to N2 animals. 24

Avg. % α- Incorporation Citrate ketoglutarate Succinate Fumarate Malate N2 1.1±0.9 3.7±2.4 3.0±1.8 1.5±1.5 3.8±2.8 pnc-1 1.1±1.1 3.8±3.0 2.7±1.5 1.3±0.9 3.2±1.7

Table 2-2: Average % glucose label incorporation in N2 and pnc-1 mutants. Shown are LC-MS measurements of observed % incorporation ± S.D. of isotopically labeled glucose into TCA cycle metabolites in N2 and pnc-1 animals.

Figure 2-7: Flow of isotopic label after pyruvate into the TCA cycle in pnc-1 mutants. The ratios of the % of isotope glucose present in lactate and citrate following normalization for prior metabolites in glycolysis in pnc-1 mutants compared to N2. . *, 0.01

Glucose storage in pnc-1 mutants

Due to glycolytic blockage and the decrease in isotopic glucose label, we next investigated glucose storage in pnc-1 mutants. We predicted that glucose was shunted and stored elsewhere when salvage NAD+ synthesis is compromised, as a result of glycolytic blockage. To shed further light on this phenomenon, we asked if there was a difference in the regulation of the 25 glucose transporter in our mutants. Interestingly, we observed a 1.5 fold increase in the C. elegans’ glucose transporter, fgt-1, mRNA levels in our mutants compared to wild-type animals

(Figure 2-8). Global metabolic profiling revealed an increase in trehalose steady state levels in pnc-1 mutants compared to wild-type (W. Wang et al., 2015). Therefore, we asked if the isotopic glucose label in pnc-1 animals was being stored as trehalose. Our data supported the increase in trehalose steady state levels in our mutants (Figure 2-9); however, there was a significant decrease in the amount of label from glucose into trehalose in pnc-1 compared to wild-type animals (Figure 2-9). As a result, this points towards the ability for organisms to sense the sensitivity for the need of glucose from their diets when cytosolic-specific NAD+ biosynthesis is compromised; however, if or where glucose is being shunted remains a mystery.

Glucose Transporter

Figure 2-8: Glucose transporter activity is up regulated in pnc-1 mutants. Relative mRNA levels of fgt-1 in pnc-1 mutants compared to N2. . **, 0.001

26

Figure 2-9: Trehalose steady state levels are increased, but fraction of total trehalose pool labeled decreases in pnc-1 mutants. (A.) Box plot of trehalose levels normalized to wild-type animals. (B). Dot-plot of the % of trehalose isotopically labeled in N2 and pnc-1 mutants.

Discussion

Compromised NAD+ biosynthesis leads to disrupted glycolysis

Our lab previously reported that loss of salvage NAD+ synthesis via mutation to the nicotinamidase, pnc-1, led to reproductive developmental defects in C. elegans (T L Vrablik et al., 2009; Tracy L Vrablik et al., 2011). Global metabolic profiling revealed glycolysis was compromised in pnc-1 mutants. We were further able to link this compromised glycolysis to the development of the gonad being delayed in pnc-1 mutants (W. Wang et al., 2015). However, although glycolysis was compromised, functions of the mitochondria were not perturbed in pnc-

1 mutants. Taken together, this led us to hypothesize that loss of cytosolicic specific NAD+ salvage synthesis compromised glycolysis impairing reproductive development in C. elegans. 27

We decided to use the powerful tool of metabolic isotopic tracing to investigate our compromised glycolysis model in pnc-1 mutants. After a short four-hour exposure with

13 universally labeled glucose ( C6-Glucose), we were able to detect an isotopically labeled glucose pool in both wild-type animals and pnc-1 mutants. The proportion of the total glucose pool that was isotopically labeled was significantly decreased in pnc-1 mutants compared to wild-type animals. Therefore, this led us to measure the flow of isotopic label from glucose through the glycolytic steps by normalizing each subsequent metabolite to the metabolite before it in each step beginning with glucose. Once we observed the ratio of the change in isotopic flow between pnc-1 mutants and wild-type animals, we observed an increase of isotope label from glucose in DHAP relative to G6P and a significant decrease in 3PGA relative to DHAP. Based the on results, this supported our hypothesis that glycolysis is blocked at the NAD+-dependent step in loss of salvage synthesis mutants. Although glycolysis is compromised, it was interesting to observe no change in the isotopic flow of label from pyruvate to lactate in pnc-1 mutants. In the cytosol, lactate dehydrogenase oxidizes NADH back to NAD+ producing lactate from pyruvate. This hints towards a compensatory relationship occurring to maintain NAD+ homeostasis in pnc-1 mutants, suggesting that NAD is being oxidized elsewhere and has to be reduced via the pyruvate to lactate mechanism. These observations give us key insight into the ability of NAD+ biosynthesis mutants to sense their environment and control homeostatic parameters.

Requirement to maintain a supply of carbon for oxidative phosphorylation by the mitochondria

The crucial role of NAD+ in regulating the intracellular redox state has triggered reinvigorated interest in understanding the dynamics behind NAD+ metabolism and biosynthesis.

The NADH equivalents generated via glycolysis in the cytosol fuel critical processes in the 28 mitochondria. Although we were surprised to find equal amounts of pyruvate steady state levels, along with an intact TCA cycle and mitochondrial functions in pnc-1 mutants, it supports the requirement to maintain a supply of carbon for oxidative phosphorylation by the mitochondria.

We were able to detect an increase isotopic label from pyruvate entering into the TCA cycle and a steady isotopic flow rate in our loss of salvage NAD+ synthesis mutants. This supports our previous observations of consistent pyruvate and TCA cycle metabolite steady state levels pnc-1 mutants compared to wild-type animals. We originally hypothesized that an increase in amino acid catabolism was occurring to maintain a supply of carbon for oxidative phosphorylation by the mitochondria. Amino acids can be converted to pyruvate via protein degradation. Over the years, I acquired supportive preliminary evidence that supported excessive use of amino acids as an energy source to compensate for insufficient glycolytic flux in pnc-1 mutants. I observed an increase of α-ketoglutarate and glutamate steady state levels, two key metabolites involved in amino acid catabolism. Alanine aminotransferases, agxt-1 and c32F10.8, transcript levels were up-regulated in pnc-1 mutants compared to controls. Therefore, we predicted that knockdown of the alanine aminotransferase, agxt-1, would decrease pyruvate levels in pnc-1 mutants. However, pyruvate levels did not significantly decrease in the RNAi animals compared to controls. Also, we predicted to see an increase in isotope label from alanine getting to pyruvate if amino acids were the energy source of pyruvate production in our loss of salvage NAD+ synthesis mutants.

However, metabolic carbon tracing of alanine to pyruvate was inclusive. After a short five-hour exposure with isotopic alanine, I detected less isotope label from alanine being incorporated into pyruvate in our pnc-1 mutants compared to wild-type animals. This data will be further discussed in Appendix. We hypothesize that unknown compensatory mechanisms are occurring to maintain a supply of carbon to the mitochondria. 29

Disruption of NAD+ metabolism leading to developmental delay can model tumor growth and progression

We originally sought to understand how and why a lack of NAD salvage synthesis disturbed normal progression of gonad development in C. elegans. In that process, we linked this developmental impairment to glycolysis being disrupted. Modeling the response to disruptions in

NAD+ homeostasis proves to be similar to what occurs during tumor formation (Figure 2-10).

NAD+

Glycolysis

Salvage Synthesis

Pyruvate Metabolic

perturbation

How/Why ?

Reproductive Development

Figure 2-10: Compromised glycolysis due to disruption in NAD+ salvage synthesis models tumor growth and progression. Schematic illustrating how disruptions to metabolic leads to disease progression by comparing to reproductive development defect observed in C. elegans.

30

Perturbations to NAD+ salvage synthesis led to glycolysis being blocked at the NAD+ dependent step. This occurrence was further linked to the impairment of reproductive development. Although glycolysis is blocked in this model, we observed normal production of pyruvate. We can hypothesize the triggering of a metabolic alteration to happen, in order to compensate for lack of glycolytic flux. Similar metabolic response mechanisms occur during tumor growth and progression, leading to alterations in metabolism. During C. elegans’ transition into adulthood, a higher rate of energy is required for the development of the reproductive organs (A. Antebi, Yeh, Tait, Hedgecock, & Riddle, 2000; a Antebi, Culotti, &

Hedgecock, 1998; Gerisch, Weitzel, Kober-Eisermann, Rottiers, & Antebi, 2001). Functions of the mitochondria are preserved in this model, whereas reproductive development suffers. This models the competition between healthy and cancerous cells during formation of tumors.

Further elucidation into the mechanisms behind NAD+ homeostasis will provide evidence and insight into anti-cancer therapeutics.

31

Materials and Methods

C. elegans Culture and Strains C. elegans strains were maintained under standard conditions at 20° C (S. Brenner, 1974) with E. Coli OP50 or UV-irradiated OP50 serving as the food source. N2 is the reference control strain. UV-irradiated OP50 plates were prepared by GS Gene Linker UV Chamber (BioRad,

Hercules, CA) for 999 seconds (T. L. T. L. Vrablik, Huang, Lange, & Hanna-Rose, 2009; Tracy

L Vrablik et al., 2011). Complete killing of the E. coli was confirmed by absence of growth on

LB agar after incubating overnight at 37° C. The following strain and allele was used: pnc-

1(pk9605) (T. L. T. L. Vrablik et al., 2009). Strains were obtained from the CGC.

Targeted Metabolomics We performed targeted LC-MS metabolomics analysis with the Metabolomics Core

Facility at Penn State. ~50 µL of worms were collected in ddH2O, flash frozen in liquid nitrogen and stored at -80° C. 15 mL samples were extracted in 1 µL of 3:3:2 acetonitrile:isopropanol:H2O with 1 mM chlorpropamide as internal standard. Samples were homogenized using a Precellys™ 24 homogenizer. Extracts from samples were dried under vacuum, resuspended in HPLC Optima Water (Thermo Scientific, Waltham, MA) and divided into two fractions, one for LC-MS and one for BCA protein analysis. Samples were analyzed by

LC-MS using a modified version of an ion pairing reversed phase negative ion electrospray ionization method (Lu et al., 2010). Samples were separated on a Supelco (Bellefonte, PA) Titan

C18 column (100 x 2.1 mm 1.9 µm particle size) using a water-methanol gradient with tributylamine added to the aqueous mobile phase. The LC-MS platform consisted of a Dionex

Ultimate 3000 quaternary HPLC pump, a Dionex 3000 column compartment, a Dionex 3000 autosampler, and an Exactive plus orbitrap mass spectrometer controlled by Xcalibur 2.2 32 software (all from ThermoFisher Scientific, San Jose, CA). The HPLC column was maintained at

30°C and a flow rate of 200 uL/min. Solvent A was 3% aqueous methanol with 10 mM tributylamine and 15 mM acetic acid; solvent B was methanol. The gradient was 0 min., 0% B; 5 min., 20% B; 7.5 min., 20% B; 13 min., 55% B; 15.5 min., 95% B, 18.5 min., 95% B; 19 min.,

0% B; 25 min 0% B. The orbitrap was operated in negative ion mode at maximum resolution

(140,000) and scanned from m/z 85 to m/z 1000. Metabolite levels were corrected to protein concentrations determined by BCA assay (Thermo Fisher).

Metabolic Tracing with Stable Isotopes 13 Stable isotope C6-Glucose (Santa Cruz Biochemicals, Dallas, TX) was used as the metabolic tracer. To collect isotopic Trp treated C. elegans, mixed stage worms were plated on

UV-killed OP50 plates and incubate at 20° C for 72 hours. Worms were then collected with M9 solution and pelleted. To the pellet, we added 1 mL concentrated heat-killed OP50 culture, 100

µL of 200 mM isotopic glucose and M9 to a final volume of 2 mL. Liquid culture solutions were incubated at room temperature for 4 hours with gentle rocking. Worms and heat-killed OP50 were separately collected by centrifuging and washed with 15 mL autoclaved water for three times. Approximately 30-40 µL worm pellet was obtained for each sample. Targeted LC-MS metabolomics analysis was performed to measure isotope incorporation. Stable isotope d5-

Tryptophan (100 mM) (Santa Cruz Biochemicals, Dallas, TX) was used as the metabolic tracer in the pnc-1 control experiment.

Quantitative Real-Time Polymerase Chain Reaction (qRT-PCR) RNA was extracted from wild-type N2 and pnc-1 mutant animals cultured on UV- irradiated OP50 plates using TRIzol Reagent (Life Technologies, Carlsbad, CA). 2 µg total 33

RNA, quantified by NanoDrop NA-1000 Spectrophotometer (NanoDrop Technologies,

Wilmington, DE), was used for reverse transcription with the High Capacity cDNA Reverse

Transcription Kit (Applied Biosystems, Foster City, CA). Three genes, cdc-42, pmp-3 and tba-1, were used as internal reference control (Hoogewijs, Houthoofd, Matthijssens, Vandesompele, &

Vanfleteren, 2008). Real-time quantitative PCR amplifications for test and reference genes were carried out using 7.5 µL of SYBR Green (PerfeCTa SYBR Green Super Mix with ROX, Quanta

Biosciences Beverly, MA), 0.6 µL of forward and reverse primer, 1.3 µL dH2O and 5 µL of diluted cDNA for each sample in a total of 15 µL. Amplification was carried out in a 7300 Real-

Time PCR System (Applied Biosystems, Foster City, CA) with initial polymerase activation at

95°C for 10 min, followed by 40 cycles of: 95° C for 15 sec denaturation, 60° C for 60 sec for primer-specific annealing and elongation. After 40 cycles, a melting curve analysis was carried out (60° C to 95° C) to verify the specificity of amplicons. The following primers were used:

Internal Reference Genes- cdc-42-F (5’-ctgctggacaggaagattacg-3’), cdc-42-R (5’- ctcggacattctcgaatgaag-3’), pmp-3-F (5’-gttcccgtgttcatcactcat-3’), pmp-3-R (5’- acaccgtcgagaagctgtaga-3’), tba-1-F (5’-gtacactccactgatctctgctgacaag-3’) and tba-1-R (5’- ctctgtacaagaggcaaacagccatg-3’). Test Genes- fgt-1-F (5-ctaagtcagttggagcataccc-3’) and fgt-1-R

(5’-aattggtgaagggatccgag-3’).

34

Chapter 3

Eukaryotic de novo NAD+ biosynthesis from tryptophan in the absence of a QPRTase homolog

Introduction

NAD+ is found in all living cells, and is an essential coenzyme, which impacts the entire metabolome (de Figueiredo, Gossmann, Ziegler, & Schuster, 2011a). NAD+ is involved in redox reactions where it carries electrons from one reaction to another and serves as a substrate for a group of enzymes called NAD+ consumers that regulate a variety of key biological processes

(Gossmann et al., 2012a; A. A. Sauve, 2008). Hence, NAD+ biosynthesis has proven to be an attractive and promising therapeutic target for influencing health-span and obesity-related phenotypes as well as tumor growth; however, NAD+ biosynthesis and homeostasis remains understudied. Exactly how NAD+ homeostasis is maintained and the biological impact of manipulating NAD+ biosynthetic pathways remains unknown in the field. Therefore, we aim to uncover these unique roles as well as the mechanisms that maintain global NAD+ homeostasis. A complete understanding of the consequences to the cell and the organism of manipulation of

NAD+ biosynthetic pathways is necessary to fully maximize the effectiveness of this target for therapeutic benefit.

A wide range of animals and yeast synthesize NAD+ via de novo synthesis from the degradation of tryptophan, via the kyneurine pathway (Ball, Yuasa, Austin, Weiser, & Hunt,

2009). Tryptophan degradation typically occurs in the nervous system and liver of most mammals (Bogan & Brenner, 2008a; Magni et al., 2004). The derivatives of the kynurenine pathway have been commonly linked to both progression and protection of neurological 35 disorders and neurodegenerative diseases. Quinolinic acid (QA) acts as an agonist to the N- methyl-D-aspartate (NMDA) glutamate receptors (Sas, Robotka, Toldi, & Vécsei, 2007b) and is characterized as a neurotoxin (Stone & Perkins, 1981). In contrast, kynurenic acid (KYNA) acts as an antagonist to a spectrum of amino acid receptors and is considered to be a neuroprotective agent (Sas et al., 2007b). The contrast between QA and KYNA and their roles in neurological disease states indicates the importance of maintaining kynurenine homeostasis for healthy and normal brain function (Schwarcz & Pellicciari, 2002). We hypothesize that the relationship between NAD+ de novo synthesis and the kynurenine pathway is critical to normal metabolic function and homeostasis.

The C. elegans’ genome encodes all the enzymes involved in the kynurenine pathway

(Figure 3-1). However, the genome lacks the critical quinolinic acid phosphoribosyltransferase

(QPRTase) homolog that converts QA into NaMN for biosynthesis of NAD+. Because C. elegans lack an apparent QPRTase homolog, it’s been assumed that this species lack active

NAD+ de novo synthesis (de Figueiredo et al., 2011a; Tracy L Vrablik et al., 2011). Because the de novo NAD+ synthesis pathway is highly conserved, we predict that organisms lacking this pathway would be unable to clear the end product of tryptophan degradation, QA, a neurotoxin

(Sas et al., 2007b; Stone & Perkins, 1981). As a result that de novo synthesis is highly conserved and the need to clear QA, we hypothesized that C. elegans may have a functional de novo NAD+ biosynthetic pathway.

We previously found that phenotypes that are dependent on NAD+ levels could be reversed upon supplementation with QA and other kynurenine metabolites, suggesting that kyneurine (KYN) pathway may contribute to NAD biosynthesis (W. Wang Thesis, 2014). I discovered that isotopic label from Trp gets incorporated into NAD+ in wild-type animals. 36

Blocking the kyneurine pathway lowers global NAD+ levels compared to controls and prevents label from Trp getting into NAD+. We identified a candidate enzyme that can use QA as a substrate for NAD+ biosynthesis. Finally, we connect kyneurine activity to fecundity in C. elegans. This evidence supports the hypothesis that NAD+ de novo synthesis is active and contributes to C. elegans’ NAD+ biosynthetic capacity and homeostasis. Uncovering this mechanism creates a novel approach for manipulating NAD+ biosynthetic pathways, which is key for the future of therapeutics.

5SQ UEP /', BGNE ,ZO LNP ), LZOV )"" IBBP 2"

/"%

Figure 3-1: Schematic of NAD+ de novo synthesis in C. elegans. Tryptophan is first converted into N-formylkynurenine (NFK) via tryptophan/indoleamine 2,3- dioxygenase (tdo-2). Next, the product from this reaction is converted to kynurenine (KYN) by arylformamidase (afmd-1). Kyn is then used as the substrate for kynurenine 3-monooxygenase (kmo-1) to produce 3-hydroxy L-kynurenine (3HK). The product is processed by kynureninase (kynu-1) to produce 3-hydroxy anthranilate (3HAA). Finally, hydroxyanthranilate 3,4-dioxygenase (haao-1) produces 2- amino-3-carboxymuconate semialdehyde, which is simultaneously degraded into QA. QA enters into the Preiss-Handler pathway when QPRTase converts it into NaMN for the biosynthesis and production of NAD+. However, the QPRTase is missing from the C. elegans genome. 37

Results

NAD+ de novo synthesis contributes to NAD+ biosynthetic capacity

The C. elegans’ genome lacks any apparent ortholog encoding QPRTase, a key enzyme required for NAD+ de novo synthesis (Gossmann et al., 2012a; Rongvaux et al., 2003; T L

Vrablik et al., 2009) (Figure 3-1). However, all enzymes required for the synthesis of QA from tryptophan are encoded in the C. elegans genome (Figure 3-1). To investigate if the kynurenine pathway contributes to NAD+ de novo biosynthesis in C. elegans, I examined the effect of loss of the pathway on global NAD+ levels. kynu-1 encodes kynureninase and is required for conversion of 3-hydroxy-L-kynurenine to 3-hydroxy-anthranilic acid. Its loss would block tryptophan catabolism, preventing formation of QA (Majewski et al., 2016). kynu-1(tm4924), a deletion allele of kynu-1, has a decrease in global NAD+ levels compared to controls (Figure 3-2). These results suggest that the kynurenine pathway contributes to NAD+ homeostasis in C. elegans perhaps by contributing to biosynthetic capacity.

Figure 3-2: Loss of kynu-1 decreases global NAD+ levels. Shown are LC-MS measurements of global NAD+ levels in N2 and kynu-1(tm4924) mutants cultured on UV-killed OP50. Boxes show the upper and lower quartile values, + indicates the mean value and lines indicate the median. Error bars indicate the maximum and minimum of the population distribution. *, 0.01

38

To directly test if de novo NAD+ synthesis from tryptophan via QA occurs in C. elegans, we used isotopically labeled metabolic tracers. After short-term supplementation of C elegans cultures with deuterium-labeled Trp, I successfully detected isotope label in the tryptophan pool

(Figure 3-3a). Furthermore, I detected deuterium label from Trp in 15% of the QA pool and in

9% of the NAD+ pool in N2 worms (Figure 3b and 4a). This demonstrates and supports flow of

Trp to NAD+ via de novo synthesis. Next, I investigated if this flow was dependent on an active kyneurine pathway. I predicted that loss of kynu-1 would block label from Trp getting incorporated into NAD+. The Trp pool was isotopically labeled in kynu-1 mutants equal to controls (Figure 3a). However, label into QA was undetectable (Figure 3b). As expected, mutation of kynu-1 decreased the efficiency of label incorporation into NAD+ 2-fold relative to controls (Figure 3-4b). We conclude that the kynurenine pathway is required for tryptophan conversion to NAD+. This supports our hypothesis that NAD+ de novo synthesis from tryptophan is actively contributing to NAD+ biosynthetic capacity in C. elegans, although the organism lacks a functional QPRTase homolog in genome.

39

A. B.

Figure 3-3: Trp pool is isotopically labeled in N2 and kynu-1 and label from Trp into QA is undetectable in kynu-1 mutants. Shown are LC-MS measurements of (A) % incorporation of d5-Trp in N2 and kynu-1 mutants. Dot-plot represents each biological replicate in wild-type (black) and kynu-1(tm4924) mutants (blue) after 4 hours of exposure to d5-Trp. (B) % Incorporation of d5-QA in wild-type. Dot-plot represents each biological replicate in wild-type (black). QA was undetectable in kynu-1(tm4924) mutants after 4 hours of exposure to d5-Trp.

A. B.

Figure 3-4: Loss of kynu-1 blocks deuterium label supplied in Trp from being incorporated into NAD+. Shown are LC-MS measurements of % incorporation of d5-NAD in wild-type and kynu-1 mutants. (A). Dot-plot of the % of NAD isotopically labeled in N2 and kynu-1 mutants. (B). Ratio of % d5-label incorporated into Trp, QA and NAD+ in kynu-1 mutants compared to N2. ***, p<0.001, calculated with Welch’s two sample t test.

40

Supplementation with NAD+ de novo precursors reverses NAD+-dependent phenotypes

We previously reported that lower NAD+ levels caused by lack of salvage synthesis associated with mutation of pnc-1 impairs reproductive development in C. elegans (W. Wang et al., 2015). NAM levels increased 19-fold and NA levels decreased 11-fold; however, NAD+ levels resulted in only a 30% reduction in pnc-1 mutants (W. Wang et al., 2015). Therefore, we hypothesized that NAD+ biosynthetic pathways could respond to maintain global NAD+ levels. If de novo NAD+ synthesis contributes to NAD+ biosynthetic capacity in C. elegans, then I predicted that this pathway would respond to loss of salvage NAD+ synthesis. In the pnc-1 mutants, I detected an increase of tdo-2, the rate-limiting step of de novo NAD+ synthesis, transcript levels (Figure 3-5), further supporting that this pathway is active and functional. If de novo NAD+ synthesis contributes to the NAD+ pool, we reasoned that an increase in available de novo precursors in combination with detected up-regulation of tdo-2 (Figure 3-5) might ameliorate pnc-1 phenotypes. To test this hypothesis, my colleague Wenqing Wang supplemented pnc-1 mutant animals with QA, Kyn and 3HAA via soaking. QA, Kyn and 3HAA

effectively rescued the gonad delay phenotype in pnc-1

mutants (Figure 3-6). Supplementation with QA also

boosted the global levels of NAD+ in pnc-1 mutants

(Figure 3-7). This data supports that providing

precursors for de novo synthesis of NAD+ can prevent

the gonad developmental delay and restore NAD+ levels

in C. elegans.

Figure 3-5: Loss of salvage NAD+ synthesis increases of tdo-2 transcript levels. qRT-PCR analysis of tdo-2 mRNA levels in pnc-1 mutants compared to N2 on UV-killed OP50. *, 0.01

Gonad Developmental Delay

Figure 3-6: QA, Kyn and 3HAA supplementation rescues gonad delay in loss of salvage NAD+ synthesis mutants. Supplementation of 20mM QA, Kyn and 3HAA to pnc-1(pk9605) mutants via soaking effectively rescues the gonad developmental defects. In histograms, error bars are S.E. **, 0.001

Figure 3-7:QA supplementation restores NAD+ levels in loss of salvage NAD+ synthesis mutants. Shown are LC-MS measurements of global NAD+ levels in pnc-1 mutants supplemented with QA (20 mM) cultured on UV-killed OP50. Dot plots show normalized NAD+ peak area. Error bars indicate the maximum and minimum of the population distribution. *, 0.01

The NAD+ deficiency in pnc-1 mutants causes a major metabolic shift that results in perturbed levels of glycolytic intermediates (W. Wang et al., 2015). Next, I investigated if QA supplementation also reversed this phenotype in pnc-1 mutants. I measured the levels of glycolytic intermediates and observed that QA supplementation successfully reversed the changes observed in pnc-1 mutants (Figure 3-8). We conclude that boosting de novo synthesis can reverse NAD+-dependent phenotypes in pnc-1 mutants.

A. B.

C.

Figure 3-8: Supplementation with QA reverses glycolytic blockage in loss of salvage NAD+ synthesis mutants. Shown are LC-MS measurements of global of G3P and DHAP (A), 2PGA and 3PGA (B) and PEP (C) levels in pnc-1 mutants supplemented with QA (20 mM) cultured on UV-killed OP50. Dot plots show normalized metabolite peak area. Error bars indicate the maximum and minimum of the population distribution. *, 0.01

43

UMPS-1 is required for QA label to be incorporated into NAD+ biosynthesis If de novo synthesis is indeed active in C. elegans, then what is functioning in place of the key QPRTase enzyme? We hypothesized that another phosphoryribosyl transferase encoded in the genome would be required to synthesize NAD+ from QA. The C. elegans genome contains

7 annotated phosphoribosyltransferases. Interestingly, only UMPS-1 (uridine monophosphate synthetase) is a dual domain PRTase/Carboxylase, similar to QPRTase. We specifically asked if umps-1 is functionally involved in NAD+ biosynthesis by determining if it is required for QA supplementation to rescue pnc-1 phenotypes or for incorporation of label from Trp into NAD+.

As noted above, QA supplementation rescues gonad developmental delay in pnc-1 mutants. We supplemented umps-1; pnc-1 double mutants with QA and examined gonad development. We predicted that if umps-1 played a role in de novo NAD+ synthesis, then loss of umps-1 would block QA rescue of gonad delay in pnc-1 mutants. As expected, umps-1 blocked the ability of

QA to rescue the penetrant gonad delay phenotype in pnc-1 mutants (Figure 3-9). Next, I predicted that loss of umps-1 would lower NAD+ steady state levels if it were participating in

NAD+ biosynthetic capacity. Moreover, global NAD+ levels are decreased in umps-1 mutants compared to controls (Figure 3-10), similar to the decreased observed in kynu-1 mutants.

Figure 3-9: umps-1 blocks QA ability to rescue gonad delay. L4 pnc-1(pK9605) and umps-1;pnc-1 animals were scored for gonad developmental delay. **, 0.001

44

Figure 3-10: Loss of umps-1 decreases global NAD+ levels. Shown are LC-MS measurements of global NAD+ levels in N2 and umps-1(ok2703) mutants cultured on UV-killed OP50. Boxes show the upper and lower quartile values, + indicates the mean value and lines indicate the median. Error bars indicate the maximum and minimum of the population distribution. **, 0.001

If UMPS-1 substitutes as the missing QPRTase, we predict that it would block incorporation of deuterium label into NAD+. We used deuterium metabolic tracer analysis. We exposed control animals and umps-1(ok2703) mutants to deuterium labeled tryptophan for 4 hours (Figure 3-11), and measured incorporation of deuterium label into NAD+. As expected, umps-1(ok2703) decreased the proportion of the NAD+ pool that became labeled (Figure 3-12).

In contrast to loss of kynu-1, umps-1 does not block label incorporation into QA (Figure 3-11).

We examined three additional umps-1 alleles, zu456, tm6379 and mn160, for their ability to decrease label incorporation from tryptophan into NAD+ (Figure 3-12) and found results are consistent with the umps-1(ok2703) allele. Furthermore, metabolic carbon and nitrogen tracing revealed that umps-1(ok2703) blocked the incorporation of label from QA into NAD+ (Figure 3-

13). This metabolic tracing analysis provides key evidence that de novo NAD+ synthesis from 45 tryptophan contributes to NAD+ biosynthesis and that UMPS-1 is required for de novo synthesis.

This further suggests that UMPS-1 can substitute as the QPRTase missing from the C. elegans genome. This data highlights conservation of de novo NAD+ biosynthesis and demonstrates an unexpected flexibility for application of a pyrimidine biosynthesis enzyme in contributing to

NAD+ biosynthesis.

A.

B.

Figure 3-11: Trp pool is isotopically labeled in N2 and umps-1 mutants and isotope label from Trp is incorporated into QA in both N2 and umps-1 mutants. Shown are LC-MS measurements of (A) % incorporation of d5-Trp in N2 and umps-1 alleles (ok2703, zu456, tm6379 and mn160). Dot-plot represents each biological replicate in wild-type and umps-1 alleles after 4 hours of exposure to d5-Trp. (B) % Incorporation of d5-QA in wild-type and umps- 1(ok2703) mutants. Dot-plot represents each biological replicate in wild-type (black) and umps-1(ok2703) (purple) mutants after 4 hours of exposure to d5-Trp.

46

A.

B.

Figure 3-14: Loss of umps-1 blocks label in Trp from being incorporated into NAD+. Shown are LC-MS measurements of % incorporation of d5-NAD in wild-type and umps-1 alleles (ok2703, zu456, tm6379 and mn160). (A). Dot-plot of the % of NAD isotopically labeled in N2 and umps-1 alleles (ok2703, zu456, tm6379 and mn160). (B). Ratio of % d5-label incorporated into Trp, QA and NAD+ in umps-1(ok2703) mutants compared to N2. **, 0.001

47

Figure 3-14: Loss of umps-1 blocks QA incorporation into NAD+. Shown is ratio of % label from QA(13C315N) incorporated into QA and NAD+ in umps- 1(ok2703) mutants compared to N2. Three biological replicates are represented in graph.

Loss of kyneurine pathway affects reproductive development

We previously reported that NAD+ salvage synthesis is required for normal progression of reproductive development in C. elegans. I next asked if NAD+ de novo synthesis from tryptophan was also necessary for either fecundity or reproductive development. I observed a modest decrease in progeny production in kynu-1 mutants compared to controls (Figure 3-16).

Supplementation with NAD+ de novo synthesis precursor, QA, restored the brood size in kynu-1 mutants supporting the role NAD+ biosynthesis plays in fecundity (Figure 3-16). Consistent with model, this data suggests that NAD+ de novo synthesis is required for normal fecundity.

Figure 3-16: Loss of NAD+ de novo synthesis disrupts fecundity. Progeny production was recorded in wild type and kynu-1(tm4924)mutants and kynu-1 mutants supplemented with 20 mM QA. In histograms, error bars are S.D. *, 0.01

48

Discussion

Intact de novo NAD+ biosynthesis in the absence of QPRTase homolog

NAD+ metabolism is at the core of critical biological processes. Thus, cells use more than one biosynthetic route for the production of NAD+ (de Figueiredo et al., 2011a). All of the pathways that contribute to NAD+ biosynthetic capacity are also highly conserved throughout evolution, alluding the importance of NAD+ as the cellular hub for metabolism in all organisms

(de Figueiredo et al., 2011a; Gossmann et al., 2012b; A. A. Sauve, 2008). NAD+ biosynthesis is critical for normal and healthy metabolic function for organisms.

In support of the hypothesis that NAD+ de novo synthesis is actively contributing to biosynthetic capacity, blocking this pathway led to a global decrease in NAD+ levels. Applying stable isotope metabolic tracer analysis, we were able to observe a decrease of incorporation from deuterium labeled tryptophan into NAD+ in mutants lacking functional de novo synthesis.

This supports the conclusion that de novo synthesis from the essential amino acid tryptophan is actively contributing to NAD+ biosynthetic capacity in C. elegans.

Using both genetics and metabolic tracer approaches, we investigated the role of UMPS-

1 in NAD+ de novo synthesis. QA supplementation was unable to rescue gonad delay in umps-

1;pnc-1 double mutants, as expected if UMPS-1 is responsible for the conversion QA into

NaMN. Loss of umps-1 decreases global NAD+ levels and incorporation from deuterium labeled tryptophan into NAD+, supporting the role of UMPS-1 functioning as the missing QPRTase enzyme in C. elegans. This supports the new activity for an old enzyme, UMPS-1, illustrating underground metabolism within NAD+ biosynthesis.

49

Requirement for NAD+ de novo biosynthesis for normal reproductive development

We previously reported the requirement of salvage NAD+ synthesis for the normal progression of gonad development and glucose metabolism in the cytosol of C. elegans.

Interestingly, loss of NAD+ de novo synthesis also resulted in decreased progeny production compared to wild type. Supplementations with NAD+ precursors were able to reverse the brood size defect in kynu-1 mutants, supporting the importance of NAD+ biosynthesis for normal reproduction development function. Both de novo NAD+ synthesis and salvage NAD+ synthesis

(Huang & Hanna-Rose, 2006) are involved in fecundity. This highlights the role of maintaining global NAD+ biosynthetic capacity for reproductive development.

50

Materials and Methods

C. elegans Culture and Strains C. elegans strains were maintained under standard conditions at 20° C (S. Brenner, 1974) with E. Coli OP50 or UV-irradiated OP50 serving as the food source. N2 is the reference control strain. UV-irradiated OP50 plates were prepared by GS Gene Linker UV Chamber (BioRad,

Hercules, CA) for 999 seconds (T. L. T. L. Vrablik et al., 2009; Tracy L Vrablik et al., 2011).

Complete killing of the E. coli was confirmed by absence of growth on LB agar after incubating overnight at 37° C. The following strains and alleles were used: pnc-1(pk9605) (T. L. T. L.

Vrablik et al., 2009), kynu-1(tm4924) and umps-1(ok2703, mn160, tm6379 and zu456). Allele umps-1(ok2703) deletes portion of the N-terminus of neighboring gene, spp-1. Strains were obtained from the CGC and Mitani Lab/National BioResource Project, Japan.

Metabolite Supplementation Nicotinic Acid (NA, Alfa Aesar, Tewksbury, MA) and Quinolinic acid (QA, MP

Biomedicals, Santa Ana, CA) supplementation were performed on culture plates. We added filter sterilized 25 mM stock solution of NA and QA to UV-irradiated plates and incubated plates at room temperature for 2 to 3 days to allow chemicals to diffuse before use.

QA, Kynurenine (Kyn, Sigma-Aldrich, St. Louis, MO) and 3-hydroxy anthranilate

(HAA, Sigma-Aldrich, St. Louis, MO) supplementation was performed in small-volume liquid culture because of limited availability of supplements. QA supplementation was also performed in liquid culture as a control for consistency between experiments. We first plated synchronized

L1 animals on UV-irradiated OP50 plates for 24 hours. 2-3 plates of synchronized L3 animals were then collected with M9 solution and pelleted. To the pellet, we added 5 µL of concentrated heat-killed OP50 culture, supplement stock solution diluted to the experimental concentration 51 and M9 to a final volume of 100 µL. Stock solutions were as follows: 20 mM QA, 20 mM Kyn or 20 mM HAA. Liquid culture solutions were incubated at room temperature for 48 hours with gentle rocking. Finally animals were plated on UV-irradiated OP50 plates and gonad development was scored when animals reached mid-L4 stage.

Phenotypic Analysis Gonad Developmental Delay: Gonad developmental delay phenotype was scored as previously reported (T L Vrablik et al., 2009). Briefly, mid-L4 stage animals with an open lumen in both the vulva and the uterus were reported as normal. “Delayed” animals are those that do not yet have an open uterine lumen when the vulva lumen achieves its characteristic mid-L4 stage morphology. We plated synchronized L1 animals on NGM plates of targeted condition and scored gonad delay when they reached mid L4 stage and calculated percentage of normal animals.

Brood Size: Young L3 stage animals were individually plated and production of progeny was counted for 4 days after reaching adulthood.

Targeted Metabolomics We performed targeted LC-MS metabolomics analysis with the Metabolomics Core

Facility at Penn State. ~50 µL of worms were collected in ddH2O, flash frozen in liquid nitrogen and stored at -80° C. 15 mL samples were extracted in 1 mL of 3:3:2 acetonitrile:isopropanol:H2O with 1 mM chlorpropamide as internal standard. Samples were homogenized using a Precellys™ 24 homogenizer. Extracts from samples were dried under vacuum, resuspended in HPLC Optima Water (Thermo Scientific, Waltham, MA) and divided into two fractions, one for LC-MS and one for BCA protein analysis. Samples were analyzed by 52

LC-MS using a modified version of an ion pairing reversed phase negative ion electrospray ionization method (Lu et al., 2010). Samples were separated on a Supelco (Bellefonte, PA) Titan

C18 column (100 x 2.1 mm 1.9 µm particle size) using a water-methanol gradient with tributylamine added to the aqueous mobile phase. The LC-MS platform consisted of a Dionex

Ultimate 3000 quaternary HPLC pump, a Dionex 3000 column compartment, a Dionex 3000 autosampler, and an Exactive plus orbitrap mass spectrometer controlled by Xcalibur 2.2 software (all from ThermoFisher Scientific, San Jose, CA). The HPLC column was maintained at

30°C and a flow rate of 200 uL/min. Solvent A was 3% aqueous methanol with 10 mM tributylamine and 15 mM acetic acid; solvent B was methanol. The gradient was 0 min., 0% B; 5 min., 20% B; 7.5 min., 20% B; 13 min., 55% B; 15.5 min., 95% B, 18.5 min., 95% B; 19 min.,

0% B; 25 min 0% B. The orbitrap was operated in negative ion mode at maximum resolution

(140,000) and scanned from m/z 85 to m/z 1000. Metabolite levels were corrected to protein concentrations determined by BCA assay (Thermo Fisher).

Metabolic Tracing with Stable Isotopes Stable isotope d5-Tryptophan (Santa Cruz Biochemicals, Dallas, TX) was used as the metabolic tracer. To collect isotopic Trp treated C. elegans, mixed stage worms were plated on

UV-killed OP50 plates and incubate at 20° C for 72 hours. Worms were then collected with M9 solution and pelleted. To the pellet, we added 1 mL concentrated heat-killed OP50 culture, 100

µL of 100 mM isotopic Trp and M9 to a final volume of 2 mL. Liquid culture solutions were incubated at room temperature for 4 hours with gentle rocking. Worms and heat-killed OP50 were separately collected by centrifuging and washed with 15 mL autoclaved water for three times. Approximately 30-40 µL worm pellet was obtained for each sample. Targeted LC-MS metabolomics analysis was performed to measure isotope incorporation. 53

Quantitative Real-Time Polymerase Chain Reaction (qRT-PCR) RNA was extracted from wild-type N2 and pnc-1 mutant animals cultured on UV- irradiated OP50 plates using TRIzol Reagent (Life Technologies, Carlsbad, CA). 2 µg total

RNA, quantified by NanoDrop NA-1000 Spectrophotometer (NanoDrop Technologies,

Wilmington, DE), was used for reverse transcription with the High Capacity cDNA Reverse

Transcription Kit (Applied Biosystems, Foster City, CA). Three genes, cdc-42, pmp-3 and tba-1, were used as internal reference control (Hoogewijs et al., 2008). Real-time quantitative PCR amplifications for test and reference genes were carried out using 7.5 µL of SYBR Green

(PerfeCTa SYBR Green Super Mix with ROX, Quanta Biosciences Beverly, MA), 0.6 µL of forward and reverse primer, 1.3 µL dH2O and 5 µL of diluted cDNA for each sample in a total of

15 µL. Amplification was carried out in a 7300 Real-Time PCR System (Applied Biosystems,

Foster City, CA) with initial polymerase activation at 95°C for 10 min, followed by 40 cycles of:

95° C for 15 sec denaturation, 60° C for 60 sec for primer-specific annealing and elongation.

After 40 cycles, a melting curve analysis was carried out (60° C to 95° C) to verify the specificity of amplicons. The following primers were used: Internal Reference Genes- cdc-42-F

(5’-ctgctggacaggaagattacg-3’), cdc-42-R (5’-ctcggacattctcgaatgaag-3’), pmp-3-F (5’- gttcccgtgttcatcactcat-3’), pmp-3-R (5’-acaccgtcgagaagctgtaga-3’), tba-1-F (5’- gtacactccactgatctctgctgacaag-3’) and tba-1-R (5’-ctctgtacaagaggcaaacagccatg-3’). Test Genes- tdo-2-F (5-tgtccgtatttgggttctgg-3’) and tdo-2-R (5’-accaactaacctgtagatattcggaa-3’).

54

Chapter 4

Nicotinamide Riboside contributes to NAD+ biosynthesis and embryonic hatching in C. elegans

This chapter has been adapted with permission from:

Wang, W., McReynolds, M. R., Goncalves, J. F., Shu, M., Dhondt, I., Braeckman, B. P., … Hanna-rose, X. W. (2015). “Comparative Metabolomic Profiling Reveals That Dysregulated Glycolysis Stemming from Lack of Salvage NAD+ Biosynthesis Impairs Reproductive Development in Caenorhabditis elegans” JBC 290(43), 26163–26179.

I have extracted my work from the original publication and present it as a separate story.

Introduction

Nicotinamide riboside as a precursor for NAD+ biosynthesis

+ People experiencing a deficiency of NAD precursors, from Vitamin B3, in their diets can develop pellagra (Belenky et al., 2007). This potentially fatal disease is characterized by dermatitis, diarrhea, dementia and death (Hegyi et al., 2004). Onset of pellagra originally pointed

+ + towards the importance of having vitamin B3 and NAD precursors in our diets. NAD homeostasis is not only important for the prevention of pellagra, recent research suggests NAD+ association with life and health span benefits (Pirinen et al., 2014; A. A. Sauve, 2008).

Nicotinamide riboside (NR) was recently discovered to be an additional vitamin B3 precursor for

NAD+ biosynthesis (Bogan & Brenner, 2008a). This salvageable precursor of NAD+ is found naturally in cow’s milk (Bieganowski & Brenner, 2004). NR has great potential as a vitamin supplement able to elevate or maintain NAD+ in specific tissues, and there has been a big push by researchers to investigate NR as a therapeutic agent. Specifically, recent studies have shown the NAD+ precursor, NR, protects against metabolic disease (Cantó et al., 2012; Gariani et al.,

2016; Lee, Hong, Jun, & Yang, 2015), neurodegenerative disorders (Gong et al., 2013) and age- 55 related decline (Mills et al., 2016) in mammals. NR supplementation allows mice to resist weight gain on a high-fat diet (Cantó et al., 2012). Notably, NR prevents noise-induced hearing loss in mice (C. Brenner, 2014; Brown et al., 2014). NR also maintains the regenerative potential of stem cells in mice that are aging (Gariani et al., 2016). Although the beneficial factors of NR supplementation are becoming known, the developmental and physiological role of NR synthesis remains to be fully elucidated.

NR and nicotinic acid riboside (NaR) can be converted to NAD+ by the NMRK pathway via either a two-step (Bieganowski & Brenner, 2004) or three-step pathway (Belenky et al.,

2007) (Figure 4-1). Nicotinamide riboside kinase 1 (NRK1) is the rate-limiting factor for the use of exogenous NR for NAD+ synthesis (Ratajczak et al., 2016). Although NMRK is a highly conserved enzyme, there was not a homolog of NMRK annotated in the C. elegans genome.

Using BLAST, we identified a candidate gene, T27A3.6, and hypothesized that this gene encodes NMRK activity. In this chapter, I show that NR contributes to NAD+ biosynthetic capacity via the NMRK pathway in C. elegans. Supplementation with NR reverses NAD+- dependent phenotypes (W. Wang et al., 2015). We also show the ability of the NMRK gene to respond to the loss of salvage NAD+ synthesis, hinting towards homeostatic interactions.

Interestingly, we discovered a novel key link between the NMRK pathway and embryonic

hatching during development. This work lays a

solid foundation to understanding the /3 /B3 developmental and physiological contributions of

NR to NAD+ homeostasis. /./ /B./ Figure 4-1: NMRK-mediated synthesis for NAD+ biosynthetic capacity. Schematic of NR being converted to NAD+ via the two-step pathway (pink), and NaR being converted to /"% NAD+ via the three-step pathway (green). 56

Results

Supplementation with NR reverses NAD+-dependent phenotypes

The nicotinamidase, pnc-1, is responsible for converting NAM into NA for NAD+ recycling via salvage synthesis, and pnc-1 mutants have a reproductive development delay that results directly from NAD+ depletion (W. Wang et al., 2015). We hypothesized that supplementation of pnc-1 mutants with NAD+ precursors that can be used to produce NAD+ via other routes could reverse defects associated with NAD+ depletion. My colleague, Wenqing

Wang, previously showed that the pnc-1 gonad delay phenotype is rescued by providing alternative NAD+ precursors including NR (W. Wang et al., 2015). I next asked if supplementation with NR could reverse other phenotypes associated with loss of salvage NAD+ synthesis. First, I supplemented C. elegans cultures with 1.25 mM NR and performed targeted metabolomics to measure the capacity of boosting NMRK-mediated synthesis to reverse metabolic perturbations in pnc-1 mutants. NR supplementation restored NAD+ levels in pnc-1 mutants, similar to providing NA as a supplement (Figure 4-2). Furthering supporting the ability of NR to reverse NAD+-dependent phenotypes, NR supplementation restored the levels of

glycolytic intermediates in pnc-1 mutants (Figure

4-3). This data suggests that NR is efficiently

used as a NAD+ precursor in C. elegans.

Figure 4-2: Both NR and NA supplementation restore NAD+ in pnc-1 mutants. Shown are LC-MS measurements of global NAD+ levels in pnc-1 mutants supplemented with both NAD+ precursors NA (25 mM) and NR (1.25 mM) cultured on UV-killed OP50. Boxes show the upper and lower quartile values, + indicates the mean value and lines indicate the median. Error bars indicate the maximum and minimum of the population distribution. *, 0.01

A. B.

D. (MVDPTF C. (1

' 1

(1 %)"1 /"% /"%)  #1(

1("

1&1

1ZSVWBUF

Figure 4-3: Both NR and NA supplementation reverses pnc-1-mediated changes in levels of glycolytic intermediates. Shown are LC-MS measurements of G3P and DHAP (A), 2PGA and 3PGA (B) and PEP (C) in pnc-1 mutants supplemented with 25 mM NA or 1.25 mM NR cultured on UV-killed OP50. Schematic indicating proposed Glycolytic blockage in pnc-1 mutants (D). Two pairs of metabolites, G3P and DHAP (A) and 2PGA and 3PGA (B) have identical molecular masses and were not separated chromatographically in this experiment, and thus the relative levels determined by LC-MS represent the total amount of both metabolites. Boxes show the upper and lower quartile values, + indicates the mean value and lines indicate the median. Error bars indicate the maximum and minimum of the population distribution. *, 0.01

Nicotinamide riboside kinase (NMRK) converts NR to NMN for NAD+ biosynthesis.

NMRK was not previously annotated in the C. elegans genome, although it’s a highly conserved enzyme. However, we identified a candidate gene, T27A3.6, using BLAST. Wenqing also provided evidence that showed loss of T27A3.6 blocked the ability of NR to rescue the reproductive gonad developmental delay associated with loss of salvage NAD+ synthesis in pnc-

1 mutants. We hypothesize that the C. elegans’ gene T27A3.6 encodes a nicotinamide riboside kinase, and I have renamed the gene nmrk-1. This work explores the dynamic function and role of NMRK-mediated synthesis in C. elegans.

NR contributes to NAD+ biosynthesis

Boosting NMRK-mediated synthesis via NR supplementation reverses phenotypes associated with NAD+ depletion, suggesting that NR is an active precursor to NAD+ in C. elegans. Therefore, we hypothesize that NR is contributing to NAD+ biosynthetic capacity in C. elegans. Next, I turned to targeted metabolomics to gain an understanding regarding the contribution of NMRK-mediated synthesis to NAD+ homeostasis. I predict that loss of NMRK- mediated synthesis would lead to global NAD+ depletion if this pathway contributed to NAD+

biosynthetic capacity. As expected, loss of nmrk-1

resulted in a 50% decrease in global NAD+ levels

compared to controls (Figure 4-4). This data

supports the conclusion that NMRK-mediated

synthesis, via T27A3.6, is contributing to NAD+

biosynthetic capacity in C. elegans.

Figure 4-4: Loss of nmrk-1 decreases global NAD+ levels. Shown are LC-MS measurements of global NAD+ levels in N2 and nmrk-1 mutants cultured on UV-killed OP50. 59

Boxes show the upper and lower quartile values, + indicates the mean value and lines indicate the median. Error bars indicate the maximum and minimum of the population distribution. **, 0.001

Due to the important nature of NAD+ in redox reactions and as a signaling molecule,

+ homeostatic parameters are in place to maintain NAD biosynthesis and metabolism. Vitamin B3

precursors from our diets serve as a source of NAD+ biosynthetic precursors. We observed that

C. elegans have the ability to maintain NAD+ homeostasis from precursor metabolites found in

their bacterial diet. Phenotypes associated with NAD+ depletion become more penetrant when

worms are fed UV-killed OP50, the bacterial strain. We hypothesize that the worms are no

longer able to use precursors in the diet to maintain NAD+ homeostasis when fed UV treated

food. Interestingly, I found that nmrk-1 mRNA levels are up-regulated in pnc-1 mutants on UV-

killed OP50 and not live OP50 (Figure 4-5). This data indicates that NMRK-mediated synthesis

from NR is able to respond to loss of salvage NAD+ synthesis although NAD+ precursors from

the diet are unavailable. This data supports NR contribution to NAD+ biosynthetic capacity, and

T27A3.6 encoding the nicotinamide

riboside kinase in C. elegans.

mRNA 1

- /N2) Figure 4-5: Loss of salvage NAD+ 1 - synthesis results in up-regulated nmrk nmrk-1 mRNA levels. pnc expression ( qRT-PCR analysis of nmrk-1 mRNA levels in pnc-1 mutants compared to N2 on both live and UV-killed OP50. ***, Ratio of p<0.001, calculated with Student’s t test.

60

NR contributes to embryonic hatching during development

in utero and ex utero development in C. elegans’ embryos typically takes up to 12-14 hours. By surprise, I discovered that loss of NMRK-mediated synthesis had the ability to extend embryogenesis for an overwhelming 240 hours under UV-killed OP50 conditions (Figure 4-6).

Electron transport chain mutants are the only known condition known to extend embryogenesis dramatically. They can extend hatching time by 50% to 24-28 hours (Felkai et al., 1999; J. Feng,

Bussière, & Hekimi, 2001). In order to investigate extended embryogenesis coupled with our

UV-killed OP50 condition and to compare extension observed with nmrk-1 mutants to mutants that extend embryogenesis, I decided to observe embryonic development on the electron transport chain mutants, clk-1 and isp-1, on our UV-killed OP50 food source. Embryogenesis was extended for 28 hours in the clk-1 mutants (Figure 4-7) and for 72 hours in the isp-1 mutants

(Figure 4-8). This extension was slightly longer on UV-killed OP50 compared to live OP50.

However, three days of extended embryogenesis cannot compare to the ten days nmrk-1 mutants can extend embryonic development. The extension in clk-1 mutants was only in a small percent of the brood, compared to isp-1 mutants; whereas, the proportion of the brood that is extended in nmrk-1 mutants is greater. I conclude that this unique phenotype could be due to either a slower embryogenesis or normal embryogenesis with a leaky failure to hatch. Current data examining the progression of embryonic development over time, our undergraduate team has shown no difference in embryogenesis progression, suggesting the latter. These findings support a novel role involving NMRK-mediated synthesis and normal embryonic hatching during development in C. elegans.

61

Figure 4-6: Embryogenesis is extended for 240 hours in nmrk-1 mutants. Embryonic development was scored by counting number of embryo hatching over a period of time in N2 and nmrk-1 mutants cultured on UV-killed OP50.

Figure 4-7: Embryogenesis is extended for up to 28 hours in clk-1 mutants. Embryonic development was scored by counting number of embryo hatching over a period of time in N2 and clk-1 mutants cultured on UV-killed OP50. 62

Figure 4-8:Embryogenesis is extended for up to 72 hours in isp-1 mutants. Embryonic development was scored by counting number of embryo hatching over a period of time in N2 and isp-1 mutants cultured on UV-killed OP50.

There is a plethora of evidence in the literature supporting the role of NR in promoting healthy aging and improving age-associated aliments (Brown et al., 2014; Cantó et al., 2012;

Mills et al., 2016; Trammell et al., 2016). We next asked if there were any obvious changes between life and health-span of nmrk-1 mutants on UV-killed OP50 hatched on the first day versus the seventh day. An undergraduate I worked closely with, Sarah Chang, performed a lifespan assay. We discovered that the life and health-span of nmrk-1 mutants, hatched on both day one and seven, was comparable to wild-type animals on UV-killed OP50 (Figure 4-9).

Therefore, this supports a novel role and function for NMRK-mediated synthesis in C. elegans embryogenesis.

63

120

100

80

60 nmrk-1 DF, d7 nmrk-1 DF, d1 40 N2 DF

Percent Survival 20

0 0.00 10.00 20.00 30.00 40.00 Day of adulthood

Figure 4-9: Health and life span in nmrk-1 mutants are comparable to N2. Lifespan of N2 and nmrk-1 mutants hatched on both day 1 and day 7 cultured on UV-killed OP50 at 20° C.

Discussion

NR contribution to the cellular NAD+ pool

NR supplementation has the ability the reverse NAD+-dependent phenotypes associated with loss of salvage NAD+ synthesis. We hypothesized that NMRK-mediated synthesis contributed to NAD+ biosynthetic capacity as well in C. elegans. Targeted metabolomics revealed a 50% decrease in global NAD+ levels in nmrk-1 mutants compared to wild-type animals (Figure 4-4). Thus, NMRK-mediated synthesis contributes to the cellular NAD+ pool in

C. elegans. Furthering supporting this conclusion, I discovered a response of nmrk-1 mRNA levels in loss of salvage NAD+ synthesis mutants on UV-killed OP50 (Figure 4-5). However, this 64 observation was not observed in pnc-1 mutants on live OP50 (Figure 4-5). This hints toward the existence of homeostatic mechanisms occurring to maintain NAD+ homeostasis. Together, this furthers supports and provides the physiological dynamics of NMRK-mediated synthesis contributing to global NAD+ biosynthesis and metabolism in C. elegans.

NR contribution to C. elegans’ embryogenesis

Typically, C. elegans’ embryonic development takes between 12-14 hours in wild-type animals under normal conditions. However, embryogenesis is extended in mutants with a defective electron transport chain. Two mutants, clk-1 and isp-1, extend embryonic development for up to 24 and 48 hours respectively (Felkai et al., 1999; J. Feng et al., 2001). clk-1 encodes for a highly conserved demethoxyubiquinone (DMQ) hydroxylase ortholog, responsible for the biosynthesis of coenzyme Q from 5-demethoxyubiquinone (Felkai et al., 1999), whereas, isp-

1encodes an iron sulphur protein subunit of the mitochondrial complex III in the mitochondrial membrane (J. Feng et al., 2001). Both genes are key to the normal functions of oxidative phosphorylation, and mutation to the genes supports the onset of extended embryogenesis. To our surprise, loss of NMRK-mediated synthesis extended embryonic development for 10 days in

C. elegans when cultured on UV-killed OP50. Embryogenesis in clk-1 and isp-1 mutants was extended for up to 3 days with the UV-killed nutritional condition. This reveals a novel and dynamic role for NMRK-mediated synthesis to C. elegans’ embryogenesis further linking NAD+ biosynthesis and metabolism to reproductive development.

We also examined the life and health span of nmrk-1 mutants hatched on day 1 and day 7.

Surprisingly, we discovered that nmrk-1 mutants hatched on either day had a comparable life and health-span trend as wild-type animals on UV-killed OP50. To gain a deeper understanding of 65 the intricate details involving NMRK-mediated synthesis, UV-killed OP50 and embryonic development, we assigned this project to a team of undergraduates to elucidate and uncover the novel mechanisms involved. So far, this team has discovered that nmrk-1 embryos undergo normal embryogenesis but are unable to hatch at proper time. We hypothesize that the mutants enter a state similar to L1 arrest during the last stage of embryonic development until they either hatch or die over the 10-day period. We also have evidence in our global metabolomics analysis that suggested that culture of C. elegans on UV-killed E. coli caused changes to metabolites indicative of oxidative stress. Our team of undergraduates also uncovered a link between oxidative stress and the extended embryogenesis in nmrk-1 mutants fed UV-killed OP50.

Supplementing with paraquat, which induces ox stress, can mimic this phenotype in nmrk-1 mutants similar to UV-killed OP50. Together, this data supports a novel mechanism underlying

NAD+ biosynthesis and reproductive development for NMRK-mediated synthesis.

66

Materials and Methods

C. elegans strains and culture

C. elegans strains were maintained under standard conditions at 20° C (S. Brenner, 1974) with E. Coli OP50 or UV-irradiated OP50 serving as the food source. N2 is the reference control strain. UV-irradiated OP50 plates were prepared by GS Gene Linker UV Chamber (BioRad,

Hercules, CA) for 999 seconds (T. L. T. L. Vrablik et al., 2009; Tracy L Vrablik et al., 2011).

Complete killing of the E. coli was confirmed by absence of growth on LB agar after incubating overnight at 37° C. The following strains and alleles were used: nmrk-1(ok2571), clk-1(qm30 or e2519) and isp-1(qm150). Strains were obtained from the CGC.

Metabolite supplementation

Nicotinamide riboside (NR, CTMedChem) supplementation was performed in small- volume liquid culture instead of adding supplement to culture plates because of limited availability of the supplement. First synchronized L1 animals were plated on UV-irradiated

OP50 plates for 36 hours. 2-3 plates of synchronized L2-L3 animals were then collected with M9 solution and pelleted. To the pellet, I added 5 µL concentrated heat-killed OP50 culture

(prepared by autoclaving 100 mL of OP50 overnight culture, pelleting, removing 90 mL of supernatant and vortexing to resuspend), supplement 1 mM NR stock solution diluted to 100 µM and M9 to a final volume of 100 µL. Liquid culture solutions were incubated at room temperature for 24 hours with gentle rocking. Finally animals collected and flash frozen in liquid nitrogen to be processed for targeted metabolomics.

67

Targeted metabolomics

We performed targeted LC-MS metabolomics analysis with the Metabolomics Core

Facility at Penn State. ~50 µL of worms were collected in ddH2O, flash frozen in liquid nitrogen and stored at -80° C. 15 mL samples were extracted in 1 mL of 3:3:2 acetonitrile:isopropanol:H2O with 1 mM chlorpropamide as internal standard. Samples were homogenized using a Precellys™ 24 homogenizer. Extracts from samples were dried under vacuum, resuspended in HPLC Optima Water (Thermo Scientific, Waltham, MA) and divided into two fractions, one for LC-MS and one for BCA protein analysis. Samples were analyzed by

LC-MS using a modified version of an ion pairing reversed phase negative ion electrospray ionization method (Lu et al., 2010). Samples were separated on a Supelco (Bellefonte, PA) Titan

C18 column (100 x 2.1 mm 1.9 µm particle size) using a water-methanol gradient with tributylamine added to the aqueous mobile phase. The LC-MS platform consisted of a Dionex

Ultimate 3000 quaternary HPLC pump, a Dionex 3000 column compartment, a Dionex 3000 autosampler, and an Exactive plus orbitrap mass spectrometer controlled by Xcalibur 2.2 software (all from ThermoFisher Scientific, San Jose, CA). The HPLC column was maintained at

30°C and a flow rate of 200 uL/min. Solvent A was 3% aqueous methanol with 10 mM tributylamine and 15 mM acetic acid; solvent B was methanol. The gradient was 0 min., 0% B; 5 min., 20% B; 7.5 min., 20% B; 13 min., 55% B; 15.5 min., 95% B, 18.5 min., 95% B; 19 min.,

0% B; 25 min 0% B. The orbitrap was operated in negative ion mode at maximum resolution

(140,000) and scanned from m/z 85 to m/z 1000. Metabolite levels were corrected to protein concentrations determined by BCA assay (Thermo Fisher).

68

Phenotypic Analysis Ex utero embryonic development: Mixed staged L3/L4 N2, nmrk-1, clk-1 and isp-1 mutants were transferred and maintained on either live or UV-killed OP50. The following day,

10-15 adult worms for each strain were plated to the appropriated live or UV-killed OP50 plate and allowed to lay eggs for 4 hours. After the 4-hour time point, worms were removed from the plates and the beginning number of eggs was counted. The number of embryos hatched was accounted for an every time point, until all eggs either died or hatched. We report the % hatching of embryos post fertilization.

Lifespan assay: Lifespan was assayed and scored simultaneously for N2, nmrk-1 (day 1) and nmrk-1 (day 7) mutants grown on UV-killed OP50.

Quantitative Real-Time Polymerase Chain Reaction (qRT-PCR) RNA was extracted from wild-type N2 and pnc-1 mutant animals cultured on both live

OP50 and UV-irradiated OP50 plates using TRIzol Reagent (Life Technologies, Carlsbad, CA).

2 µg total RNA, quantified by NanoDrop NA-1000 Spectrophotometer (NanoDrop

Technologies, Wilmington, DE), was used for reverse transcription with the High Capacity cDNA Reverse Transcription Kit (Applied Biosystems, Foster City, CA). Three genes, cdc-42, pmp-3 and tba-1, were used as internal reference control (Hoogewijs et al., 2008). Real-time quantitative PCR amplifications for test and reference genes were carried out using 7.5 µL of

SYBR Green (PerfeCTa SYBR Green Super Mix with ROX, Quanta Biosciences Beverly, MA),

0.6 µL of forward and reverse primer, 1.3 µL dH2O and 5 µL of diluted cDNA for each sample in a total of 15 µL. Amplification was carried out in a 7300 Real-Time PCR System (Applied

Biosystems, Foster City, CA) with initial polymerase activation at 95°C for 10 min, followed by

40 cycles of: 95° C for 15 sec denaturation, 60° C for 60 sec for primer-specific annealing and 69 elongation. After 40 cycles, a melting curve analysis was carried out (60° C to 95° C) to verify the specificity of amplicons. The following primers were used: Internal Reference Genes- cdc-

42-F (5’-ctgctggacaggaagattacg-3’), cdc-42-R (5’-ctcggacattctcgaatgaag-3’), pmp-3-F (5’- gttcccgtgttcatcactcat-3’), pmp-3-R (5’-acaccgtcgagaagctgtaga-3’), tba-1-F (5’- gtacactccactgatctctgctgacaag-3’) and tba-1-R (5’-ctctgtacaagaggcaaacagccatg-3’). Test Genes- nmrk-1-F (5-ggatcagcttgttagtcaccc-3’) and nmrk-1-R (5’-tgcacagattccacgtagc-3’).

70

Chapter 5

Compensatory roles for NAD+ biosynthetic pathways and consumers in C. elegans

This chapter has been adapted with permission from:

Wang, W., McReynolds, M. R., Goncalves, J. F., Shu, M., Dhondt, I., Braeckman, B. P., … Hanna-rose, X. W. (2015). “Comparative Metabolomic Profiling Reveals That Dysregulated Glycolysis Stemming from Lack of Salvage NAD+ Biosynthesis Impairs Reproductive Development in Caenorhabditis elegans” JBC 290(43), 26163–26179.

I have extracted my work from the original publication and present it as a separate story.

Introduction

Critical nature of NAD+ pool in cellular metabolism

As a central metabolite, NAD+ plays a critical role in modulating overall energy homeostasis through cellular metabolism. There are four known major molecules that serve as precursors for NAD+ biosynthesis, tryptophan, nicotinic acid, nicotinamide and nicotinamide riboside. However, intermediates, such as NMN can also stimulate NAD+ biosynthesis directly.

A critical balance between NAD+ biosynthetic and consuming pathways sets the NAD+ cellular pool (Srivastava, 2016). NAD+ has the dynamic ability to respond to physiological stimuli, and levels can be modulated via both physiological processes and pharmacologically (Pirinen et al.,

2014). NAD+ pools can also behave independently. Mitochondria NAD+ pools are more stable than NAD+ pools in the cytosol in order to preserve oxidative phosphorylation (Srivastava,

2016). Also, NAD+ is not distributed evenly among subcellular compartments, and intracellular

NAD+ has a very short half-life, estimated to be 1 to 2 hours (Houtkooper, Cantó, Wanders, &

Auwerx, 2010). Due to the critical function of NAD+ as a coenzyme in a majority of metabolic pathways and substrate for key biological enzymes, we predict that limitation on NAD+ would 71 perturb metabolic efficiency. Therefore, we propose that there are compensatory mechanisms in place to maintain NAD+ homeostasis.

In the previous chapters and past research in the lab, we showed that loss of a specific

NAD+ biosynthetic pathway results in separable developmental and physiological perturbations

(T. L. T. L. Vrablik et al., 2009; Tracy L Vrablik et al., 2011; W. Wang et al., 2015). Loss of salvage NAD+ synthesis resulted in dysregulated glycolysis impairing reproductive development in C. elegans (W. Wang et al., 2015). Although glycolysis is compromised in these mutants, functions of the mitochondria are intact and not perturbed (W. Wang et al., 2015). Furthermore, we observed a response of both NMRK-mediated synthesis and de novo synthesis in pnc-1 mutants. We hypothesize that a compensatory network is in place to maintain dynamic mechanisms involved with NAD+ homeostasis. In this chapter, I show that the NAD/NADH ratio is not changed in pnc-1 mutants. Also, deletion of the main NAD+ consumer, PARP, increases global NAD+ levels in pnc-1 mutants. I observed a similar response of up-regulation of

NAD+ biosynthetic pathway genes in the absence of NMRK-mediated synthesis or de novo

NAD+ synthesis, suggesting a compensatory network. Loss of both NMRK-mediated synthesis and de novo NAD+ synthesis resulted in increased glycolytic and TCA cycle metabolites steady state levels. Finally, oxygen consumption and heat production is not perturbed in any NAD+ biosynthetic mutants. This work illustrates a dynamic compensatory mechanism and network in place to maintain global NAD+ homeostasis, due to the critical nature of NAD+ cellular pool in energy metabolism and homeostasis.

72

Results

NAD/NADH ratio is not impacted in loss of salvage NAD+ synthesis mutants

We previously reported that loss of salvage NAD+ synthesis, via mutation to the nicotinamidase pnc-1, resulted in minor depletion of NAD+ (W. Wang et al., 2015). Although the depletion to global NAD+ levels were only ~30%, glycolysis is compromised and reproductive development is impaired in pnc-1 mutants (W. Wang et al., 2015). NAD/NADH ratio and redox state is normally influenced by the breakdown and availability of dietary nutrients and energy

(Pittelli et al., n.d.; Srivastava, 2016). Therefore, I asked if the NAD/NADH ratio is perturbed in loss of salvage NAD+ synthesis mutants. I found that the NAD/NADH ratio does not change between wild-type animals and pnc-1 mutants (Figure 5-1). The NAD/NADH ratio is not affected in pnc-1 mutants, supporting the conclusion that NAD+ depletion is the source for phenotypes associated with loss of salvage NAD+ synthesis.

Figure 5-1: NAD/NADH ratio in pnc-1 mutants. [NAD]/[NADH] ratio does not differ between N2 and pnc-1 mutants under standard (p=0.8956) or UV- killed (p=0.7869) food conditions. Error bars are S.E. p values were calculated using Student’s t test.

73

PARP deletion increases NAD+ levels in loss of salvage NAD+ synthesis mutants

Supplying NAD+ biosynthesis or manipulating NAD+ consumption can directly modulate

NAD+ levels. PARP activity constitutes the main NAD+ catabolic activity, which drives cells to synthesize NAD+ (Bai et al., 2011; Chiarugi, 2012; Pirinen et al., 2014). These enzymes use

NAD+ to catalyze a reaction in which the ADP ribose moiety is transferred to a substrate protein.

PARPs are activated in response to DNA damage and genotoxic stress (Bai et al., 2011;

Chiarugi, 2012; Pirinen et al., 2014). We next investigated the consequences of deleting PARP activity in loss of salvage NAD+ synthesis mutants. Wenqing Wang provided evidence that revealed PARP deletion, via parp-1 mutation, rescued the gonad delay phenotype in pnc-1 mutants (W. Wang Thesis 2014). I predicted we could raise NAD+ levels in pnc-1 mutants via

PARP deletion. Targeted LC-MS revealed NAD+ steady state levels are increased in parp-1;pnc-

1 double mutants (Figure 5-2), supporting the notion that PARP deletion can restore NAD+

levels in loss of salvage synthesis

mutants.

Figure 5-2: PARP deletion increases global NAD+ levels in pnc-1 mutants. parp-1(ok988) loss-of-function moderately increases NAD+ levels. NAD+ levels were measured using LC-MS. Boxes show the upper and lower quartile values, + indicates the mean value and lines indicate the median. Error bars indicate the maximum and minimum of the population distribution. . *, 0.01

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Loss of NAD+ biosynthetic pathways results in a homeostatic response

Both NMRK-mediated synthesis and de novo NAD+ synthesis can respond to loss of salvage NAD+ synthesis. In the previous chapters, I showed the ability of each pathway to respond to NAD+ depletion in pnc-1 mutants. I found nmrk-1 and tdo-2 mRNA levels up- regulated in pnc-1 mutants, supporting the hypothesis that these pathways contribute to NAD+ biosynthesis in C. elegans. This observation also led me to ask if other homeostatic parameters were occurring to maintain homeostatic requirements of NAD+. First, I asked if there was a trend in nmrk-1 and pnc-1 response to loss of de novo NAD+ synthesis. In kynu-1 mutants, pnc-1 and nmrk-1 mRNA levels are up-regulated 2-fold and 8-fold (Figure 5-3). This reciprocal trend was also observed in loss of NMRK-mediated synthesis mutants. tdo-2 and pnc-1 mRNA levels are also up-regulated in nmrk-1 mutants (Figure 5-4). The ability of these pathways to respond suggests homeostatic parameters occurring amongst NAD+ biosynthetic pathways to maintain

NAD+ homeostasis.

Figure 5-3: Loss of NMRK-mediated synthesis results in up-regulated tdo-2 and pnc-1 mRNA levels. qRT-PCR analysis of tdo-2 and pnc-1 mRNA levels in nmrk-1 mutants compared to N2 on both live and UV-killed OP50. ***, p<0.001, calculated with Student’s t test.

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Figure 5-4: Loss of de novo NAD+ synthesis results in up-regulated pnc-1 and nmrk-1 mRNA levels. qRT-PCR analysis of pnc-1 and nmrk-1 mRNA levels in kynu-1 mutants compared to N2 on UV-killed OP50. ***, p<0.001, calculated with Student’s t test.

Loss of NAD+ biosynthetic pathways results in global metabolic changes

Due to the requirement for NAD+ in a majority of redox reactions, I next investigated the consequences of manipulating NAD+ biosynthesis on two major metabolic pathways, the TCA cycle and glycolysis. We expected to find metabolic perturbations in both nmrk-1 and kynu-1 mutants. Targeted LC-MS revealed perturbations in the TCA cycle at the critical steps that depend on the redox state of the NAD+ when de novo NAD+ synthesis is blocked. I observed a significant decrease in global citrate/isocitrate levels (Figure 5-5a) in kynu-1 mutants compared to wild-type animals. Global α-ketoglutarate levels exhibited an increased trend in kynu-1 mutants (Figure 5-5b), whereas succinate global levels were significantly decreased in kynu-1 mutants compared to wild-type animals (Figure 5-5c). The malate shuttle is responsible for translocating NAD+ across the mitochondrial membrane, because the mitochondrial inner 76 membrane is impermeable to NADH. These electrons then enter the electron transport chain of the mitochondria to produce ATP. Targeted metabolomics revealed a decreased trend in global malate levels in kynu-1 mutants (Figure 5-5d). Global oxaloacetate levels exhibited an increased trend in kynu-1 mutants (Figure 5-5e), suggesting blockage in the malate shuttle of kynu-1 mutants.

Figure 5-5: Loss of kynu-1 results in TCA cycle perturbations. Shown are LC-MS measurements of global TCA cycle intermediate levels in N2 and kynu-1 mutants cultured on UV-killed OP50. Boxes show the upper and lower quartile values, + indicates the mean value and lines indicate the median. Error bars indicate the maximum and minimum of the population distribution. §, 0.05

NAD+ is also required for the breakdown of glucose into pyruvate via glycolysis. We previously reported that loss of salvage NAD+ synthesis results in glycolytic blockage at the 77 sixth step that uses NAD+ to convert glyceraldehyde 3-phosphate (G3P) into 1,3- bisphosphoglycerate (1,3BPG). However, unlike pnc-1 mutants, I observed a significant increased in the global metabolite levels of the intermediates glucose 6-phosphate (G6P)/fructose

6-phosphate (F6P), dihydroxyacetone phosphate (DHAP)/G3P, 3-phosphoglycerate (3PGA)/2- phosphoglycerate (2PGA), phosphoenolpyruvate (PEP) and pyruvate involved in glycolysis in kynu-1 mutants compared to wild type (Figure 5-6). This data suggests that loss of NAD+ de novo synthesis results in increased glycolytic intermediates steady state levels.

Figure 5-6: Loss of kynu-1 results in increased glycolysis. Shown are LC-MS measurements of global glycolytic intermediate levels in N2 and kynu-1 mutants cultured on UV-killed OP50. Boxes show the upper and lower quartile values, + indicates the mean value and lines indicate the median. Error bars indicate the maximum and minimum of the population distribution. *, 0.01

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Next, I investigated the impact of loss of NMRK-mediated synthesis on the metabolic requirements in C. elegans. Interestingly, I observed increased metabolic capacity in nmrk-1 mutants. Targeted LC-MS revealed that intermediate metabolites involved in the TCA cycle were more abundant in nmrk-1 mutants compared to wild-type animals. Citrate/Isocitrate (Figure

5-7a) and succinate (Figure 5-7b) steady state levels were significantly increased, while fumarate

(Figure 5-7c) and malate (Figure 5-7d) steady state levels exhibited an upward trend. Similar to loss of de novo NAD+ synthesis mutants, nmrk-1 mutants also had increased glycolytic intermediate steady state levels. G6P/F6P, DHAP/G3P, and PEP steady state levels are increased up to 11-fold in nmrk-1 mutants; however, 3PGA steady state levels exhibited a decreased trend compared to controls (Figure 5-8). This suggests homeostatic parameters involved in maintaining requirements of NAD+ biosynthesis.

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Figure 5-7: Loss of nmrk-1 results in increased citrate/isocitrate. Shown are LC-MS measurements of global TCA cycle levels in N2 and nmrk-1 mutants cultured on UV- killed OP50. Boxes show the upper and lower quartile values, + indicates the mean value and lines indicate the median. Error bars indicate the maximum and minimum of the population distribution. *, 0.01

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Figure 5-8: Loss of nmrk-1 results in increased glycolysis. Shown are LC-MS measurements of global glycolytic intermediate levels in N2 and nmrk-1 mutants cultured on UV-killed OP50. Boxes show the upper and lower quartile values, + indicates the mean value and lines indicate the median. Error bars indicate the maximum and minimum of the population distribution. ***,

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Figure 5-9: Schematic summarizing glycolytic and TCA metabolic changes observed in loss of de novo and NMRK-mediated synthesis mutants. Blue arrows represent metabolite changes in kynu-1 mutants and purple arrows represent metabolite changes in nmrk-1 mutants. Line indicates no change in metabolite steady state levels.

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Mitochondria are protected in NAD+ biosynthetic mutants

Due to metabolic dysregulation of the TCA cycle and glycolysis in kynu-1 and nmrk-1 mutants, we decided to measure the function of the mitochondria. Oxygen consumption and heat production are two key readouts for mitochondrial function. We previously reported that both oxygen consumption and heat production were not impacted in loss of salvage NAD+ synthesis mutants (Wang 2015). Our collaborators in Bart Breckman’s lab provided this data for us. We decided to return to this collaboration, and investigate the consequences to mitochondria function in loss of de novo and NMRK-mediated synthesis mutants. We observed no change of oxygen consumption in kynu-1 and nmrk-1 mutants compared to wild type (Figure 5-10). Heat production was not changed in kynu-1 and nmrk-1 mutants (Figure 5-11). Also, mitochondria in the double and triple mutants of NAD+ biosynthetic pathways are protected. Mitochondrial functions are maintained despite perturbation to NAD+ biosynthetic pathways, suggesting a compensatory network to maintain NAD+-dependent processes.

Figure 5-10: Loss of NAD+ biosynthetic pathway does not affect oxygen consumption. Compared with N2 animals, pnc-1, nmrk-1, kynu-1, nmrk-1;pnc-1, kynu-;pnc-1 or nmrk-1;kynu-1;pnc-1 mutants do not show any significant differences in oxygen consumption as calculated by Student’s t test. Animals were cultured on standard OP50 plates. Error bars are S.E. 83

Figure 5-11: Loss of NAD+ biosynthetic pathway does not affect heat production. Compared with N2 animals, pnc-1, nmrk-1, kynu-1, nmrk-1;pnc-1, kynu-;pnc-1 or nmrk-1;kynu-1;pnc-1 mutants do not show any significant differences in heat production as calculated by Student’s t test. Animals were cultured on standard OP50 plates. Error bars are S.E.

Discussion

Compensatory network within NAD+ biosynthetic pathways and consumers to maintain global NAD+ homeostasis

Loss of salvage NAD+ synthesis compromises glycolysis and impairs reproductive development in C. elegans (W. Wang et al., 2015). Although glycolysis is blocked, the TCA cycle and functions of the mitochondria are intact. Global NAD+ levels are depleted only ~30% in pnc-1 mutants compared to wild-type animals. This observation suggests compensatory mechanisms are in place to maintain mitochondria NAD+ homeostasis. Interestingly, I discovered that the NAD/NADH ratio is not perturbed in our pnc-1 mutants. This data illustrates that manipulating salvage NAD+ synthesis directly affects the cellular NAD+ pool versus the

NAD/NADH ratio. We hypothesize that phenotypes associated with loss of a specific NAD+ biosynthetic pathway are due to direct depletion of the cellular NAD+ pool. 84

NAD+ serves as a substrate for a group of enzymes known to regulate key biological processes. PARPs are one of the main groups of enzymes that consume NAD+ for their reactions responding to DNA damage and genotoxic stress. We next investigated if deleting PARP activity could reverse NAD+-dependent phenotypes in pnc-1 mutants. Wenqing Wang provided data that revealed PARP deletion, via mutation of parp-1, could rescue the gonad delay developmental phenotype in pnc-1 mutants. I next asked if we could restore NAD+ levels in pnc-1 mutants via

PARP deletion. Targeted LC-MS revealed global NAD+ levels were increased in parp-1;pnc-1 double mutants compared to pnc-1 mutants. My lab mate, Avni Upadhyay, discovered that

PARP deletion could also reverse phenotypes associated with NAM accumulation. It would be interesting to investigate NAM steady state levels in parp-1;pnc-1 double mutants. I predict

PARP deletion would reverse NAM accumulation in pnc-1 mutants. Manipulating NAD+ consumer activity can provide homeostatic parameters in loss of salvage NAD+ synthesis mutants.

I previously showed that de novo and NMRK-mediated synthesis could respond to the lack of salvage NAD+ synthesis in C. elegans. This data originally provided evidence that supported my hypothesis that both pathways are active and contributing to NAD+ biosynthetic capacity in C. elegans. However, this data also suggested that there were homeostatic parameters in place to maintain global NAD+ homeostasis. I next asked if there was a similar trend in loss of de novo NAD+ synthesis mutants. I observed an increase in nmrk-1 and pnc-1 mRNA levels in kynu-1 mutants. Furthermore, I noticed a reciprocal trend in nmrk-1 mutants, where tdo-2 and pnc-1 mRNA levels are up-regulated. I hypothesize that there is a compensatory network amongst NAD+ biosynthetic pathways to maintain global NAD+ homeostasis. 85

Unlike pnc-1 mutants, loss of de novo NAD+ synthesis yielded metabolic perturbations in the TCA cycle at the steps that use NAD+ for the conversion of intermediates. However, glycolytic intermediates steady state levels are significantly increased, suggesting an increase glycolytic flux in kynu-1 mutants. Similar to kynu-1 mutants, nmrk-1 mutants exhibited an increase in steady state metabolite levels of TCA and glycolysis intermediates, suggesting homeostatic parameters to maintain oxidative phosphorylation. Most surprisingly, we discovered that none of the single, double or triple mutants of NAD+ biosynthesis impact oxygen consumption or heat production, the two key readouts of mitochondria function. Further analysis into this compensatory network is required to elucidate the mechanisms behind NAD+ homeostasis. This can be accomplished by elucidating the biosynthetic contribution of each pathway to the NAD+ pool in the tissues/organs in a mouse model.

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Material and Methods

C. elegans Culture and Strains

C. elegans strains were maintained under standard conditions at 20° C (S. Brenner, 1974) with E. Coli OP50 or UV-irradiated OP50 serving as the food source. N2 is the reference control strain. UV-irradiated OP50 plates were prepared by GS Gene Linker UV Chamber (BioRad,

Hercules, CA) for 999 seconds (T. L. T. L. Vrablik et al., 2009; Tracy L Vrablik et al., 2011).

Complete killing of the E. coli was confirmed by absence of growth on LB agar after incubating overnight at 37° C. The following strains and allele was used: pnc-1(pk9605) (T. L. T. L. Vrablik et al., 2009), kynu-1 (tm4924) and nmrk-1 (ok2571). Strains were obtained from the CGC and

Mitani Lab/National BioResource Project, Japan.

NAD/NADH ratio

300 µL of mixed stage N2 or pnc-1 animals (cultured on live or UV-killed OP50) were collected in M9, snap-frozen in liquid nitrogen, and stored at -80 °C. 50 µL of thawed sample was added to duplicate wells of a black 96-well plate. NAD and NADH measurements were performed using the Elite Fluorimetric NAD/NADH Ratio Assay kit (eENZYME, LLC,

Gaithersburg, MD) according to the manufacturers’ instructions.

Targeted Metabolomics

We performed targeted LC-MS metabolomics analysis with the Metabolomics Core

Facility at Penn State. ~50 mL of worms were collected in ddH2O, flash frozen in liquid nitrogen and stored at -80° C. 15 mL samples were extracted in 1 mL of 3:3:2 acetonitrile:isopropanol:H2O with 1 mM chlorpropamide as internal standard. Samples were homogenized using a Precellys™ 24 homogenizer. Extracts from samples were dried under 87 vacuum, resuspended in HPLC Optima Water (Thermo Scientific, Waltham, MA) and divided into two fractions, one for LC-MS and one for BCA protein analysis. Samples were analyzed by

LC-MS using a modified version of an ion pairing reversed phase negative ion electrospray ionization method (Lu et al., 2010). Samples were separated on a Supelco (Bellefonte, PA) Titan

C18 column (100 x 2.1 mm 1.9 µm particle size) using a water-methanol gradient with tributylamine added to the aqueous mobile phase. The LC-MS platform consisted of a Dionex

Ultimate 3000 quaternary HPLC pump, a Dionex 3000 column compartment, a Dionex 3000 autosampler, and an Exactive plus orbitrap mass spectrometer controlled by Xcalibur 2.2 software (all from ThermoFisher Scientific, San Jose, CA). The HPLC column was maintained at

30°C and a flow rate of 200 uL/min. Solvent A was 3% aqueous methanol with 10 mM tributylamine and 15 mM acetic acid; solvent B was methanol. The gradient was 0 min., 0% B; 5 min., 20% B; 7.5 min., 20% B; 13 min., 55% B; 15.5 min., 95% B, 18.5 min., 95% B; 19 min.,

0% B; 25 min 0% B. The orbitrap was operated in negative ion mode at maximum resolution

(140,000) and scanned from m/z 85 to m/z 1000. Metabolite levels were corrected to protein concentrations determined by BCA assay (Thermo Fisher).

Phenotypic Analysis Oxygen consumption and heat production: Synchronized day 2 adults were collected and washed in S-basal medium. 400–600 µL of worm suspension (dependent on worm number) was used for live oxygen consumption (928 sixchannel oxygen system, Strathkelvin Instruments) and heat production measurements (2277 Thermal Activity Monitor, Thermometric). Heat production and oxygen consumption data were normalized to total protein determined by BCA assay of a parallel aliquot of worm suspension. All essays were repeated three times independently.

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Quantitative Real-Time Polymerase Chain Reaction (qRT-PCR)

RNA was extracted from wild-type N2, kynu-1 and nmrk-1mutant animals cultured on live and UV-irradiated OP50 plates using TRIzol Reagent (Life Technologies, Carlsbad, CA). 2

µg total RNA, quantified by NanoDrop NA-1000 Spectrophotometer (NanoDrop Technologies,

Wilmington, DE), was used for reverse transcription with the High Capacity cDNA Reverse

Transcription Kit (Applied Biosystems, Foster City, CA). Three genes, cdc-42, pmp-3 and tba-1, were used as internal reference control (Hoogewijs et al., 2008). Real-time quantitative PCR amplifications for test and reference genes were carried out using 7.5 µL of SYBR Green

(PerfeCTa SYBR Green Super Mix with ROX, Quanta Biosciences Beverly, MA), 0.6 µL of forward and reverse primer, 1.3 µL dH2O and 5 µL of diluted cDNA for each sample in a total of

15 µL. Amplification was carried out in a 7300 Real-Time PCR System (Applied Biosystems,

Foster City, CA) with initial polymerase activation at 95°C for 10 min, followed by 40 cycles of:

95° C for 15 sec denaturation, 60° C for 60 sec for primer-specific annealing and elongation.

After 40 cycles, a melting curve analysis was carried out (60° C to 95° C) to verify the specificity of amplicons. The following primers were used: Internal Reference Genes- cdc-42-F

(5’-ctgctggacaggaagattacg-3’), cdc-42-R (5’-ctcggacattctcgaatgaag-3’), pmp-3-F (5’- gttcccgtgttcatcactcat-3’), pmp-3-R (5’-acaccgtcgagaagctgtaga-3’), tba-1-F (5’- gtacactccactgatctctgctgacaag-3’) and tba-1-R (5’-ctctgtacaagaggcaaacagccatg-3’). Test Genes- tdo-2-F (5-tgtccgtatttgggttctgg-3’) and tdo-2-R (5’-accaactaacctgtagatattcggaa-3’), pnc-1-F (5- -

3’) and pnc-1-R (5’- -3’) and nmrk-1-F (5-ggatcagcttgttagtcaccc-3’) and nmrk-1-R (5’- tgcacagattccacgtagc-3’).

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Chapter 6

Discussion

Developmental and physiological roles of NAD+ biosynthetic pathways

I sought to elucidate the developmental and physiological requirements of NAD+ biosynthetic pathways. I incorporated stable isotope tracing and targeted metabolomics together with genetic manipulation of NAD+ biosynthetic pathways in order to investigate metabolic perturbations and requirements of salvage NAD+ synthesis, de novo NAD+ synthesis and

NMRK-mediated synthesis in a multi-cellular organism. This work uncovers a compensatory network, diverse and unique biological roles, and suggests underground metabolic mechanisms for biosynthesis amongst NAD+ biosynthetic pathways.

Metabolic tracing reveals compromised glycolysis in pnc-1 mutants

Our lab recently proposed a model where loss of salvage NAD+ synthesis mutants had impaired reproductive development that was associated with compromised glycolysis (W. Wang et al., 2015). Using pnc-1 mutants as my model, I developed a reliable protocol tracing the flow of stable isotopes through metabolic pathways in C. elegans. Carbon tracing with universally labeled glucose demonstrated that glycolysis is indeed compromised in our pnc-1 mutants. This inhibition occurred at the NAD+-dependent step that uses glyceraldehyde 3-phosphate dehydrogenase to convert G3P into 1,3-BPG. Although glycolysis is dysregulated in pnc-1 mutants, we observed a comparable trend to controls for the steady state levels of pyruvate and

TCA cycle metabolites (W. Wang et al., 2015). Carbon tracing revealed that the flow of label to pyruvate from PEP was intact in pnc-1 mutants and controls. Moreover, an increase flow of isotopic label from pyruvate went into citrate in pnc-1 mutants. This supports our hypothesis that 90 homeostatic parameters are in place to maintain oxidative phosphorylation by providing a steady supply of carbon to the electron transport chain.

We next investigated glucose storage in pnc-1 mutants. We predicted that glucose was shunted and stored elsewhere when salvage NAD+ synthesis is compromised, as a result of glycolytic blockage. Interestingly, we noticed an up-regulation of the glucose transporter, fgt-1, mRNA levels in pnc-1 mutants compared to controls. This result was surprising and not what I expected. Because glycolysis is dysregulated in pnc-1 mutants, I predicted that the glucose transporter activity would be down-regulated. On top of that, trehalose steady state levels were increased in pnc-1 mutants (W. Wang et al., 2015). However, isotope label from glucose was not shunted towards trehalose in loss of salvage NAD+ synthesis mutants. Further investigation is required to elucidate the mechanisms controlling glucose storage when glycolysis is compromised in pnc-1 mutants.

As a result of the steady flow of carbon to pyruvate and functions of the mitochondria being intact in pnc-1 mutants (W. Wang et al., 2015), I hypothesized that amino acid catabolism was compensating for lack of glycolysis. I predicted that proteins were being degraded into amino acids due to insufficient glycolysis, and those amino acids were being converted into pyruvate. I had supportive preliminary data that supported this hypothesis. Both alanine and cysteine steady state levels were increased in pnc-1 mutants, suggesting a steady available reservoir of amino acids in place to maintain carbon metabolism. We observed an increase of α- ketoglutarate and glutamate steady state levels, two key metabolites involved in amino acid catabolism. Alanine aminotransferases, agxt-1 and c32F10.8, transcript levels were up-regulated in pnc-1 mutants compared to controls. Therefore, we predicted that knockdown of the alanine aminotransferase, agxt-1, would decrease pyruvate levels in pnc-1 mutants. However, this was 91 not the outcome in our experiment. Furthermore, we predicted that we would observe an increase of isotopic label from alanine entering pyruvate in our pnc-1 mutants. After a short five-hour exposure, we did not observe this trend. It remains to be determined if amino acids are being used to compensate for insufficient glycolytic flux in our pnc-1 mutants. First, I would optimize the protocol using alanine as the metabolic tracing. After five hours, only 3% of the alanine pool was isotopically labeled in wild-type animals. I predict if the concentration of isotope label was increased, then this would yield an increased of the alanine pool labeled. I performed the experiment for three and five hours, I would increase the time of exposure to isotope label.

Furthermore, cysteine, serine, glycine and threonine can be converted into pyruvate as well.

Performing metabolic tracing with these additional amino acids could perhaps point towards the answer. The flexibility of cellular metabolism in C. elegans when NAD+ biosynthetic pathways are manipulated suggests that there are metabolic homeostatic mechanisms in higher organisms as well. Elucidating the homeostatic responses to cellular metabolism when targeting NAD+ biosynthesis and metabolism for therapeutic benefit is critical to maximize drug target. de novo NAD+ synthesis from tryptophan in C. elegans

C. elegans lack the key QPRTase required to convert QA into NaMN (de Figueiredo,

Gossmann, Ziegler, & Schuster, 2011b; Tracy L Vrablik et al., 2011). However, all the enzymes involved in the kyneurine pathway are present in the C. elegans’ genome. We predicted that if C. elegans did not have an active de novo NAD+ synthesis pathway, then they would be unable to clear the end product, QA, a known neurotoxin. Interestingly, we also demonstrated that QA supplementation could reverse NAD+-dependent phenotypes in pnc-1 mutants. Therefore, I hypothesized that de novo NAD+ synthesis contributed to NAD+ biosynthetic capacity, despite lacking the key QPRTase. Supporting my hypothesis, loss of de novo NAD+ synthesis, via 92 mutation of kynu-1, decreased global NAD+ levels. If de novo synthesis is active in C. elegans, then we predicted that label from isotope tryptophan would get incorporated into NAD+ in wild- type animals. As expected, I detected isotopic label from tryptophan in the NAD+ pool in C. elegans. This incorporation was dependent on the activity of the kyneurine pathway, supporting the hypothesis that de novo synthesis is contributing to NAD+ biosynthesis in C. elegans.

Furthermore, loss of kyneurine activity resulted in a decrease progeny production, and QA supplementation was able to reverse the fecundity defect. This supports a role for de novo NAD+ biosynthesis in C. elegans reproduction.

On top of establishing that de novo NAD+ synthesis is functioning in C. elegans, I also uncovered an unexpected enzyme required for both labeled QA and labeled Trp to get incorporated into NAD+ illustrating underground metabolism. The predicted enzyme activities for UMPS-1 include both the decarboxylase and phosphoribosyltransferase activities required to process QA into NAD+. This suggests it can use QA as a substrate. We predicted that if UMPS-

1 were required for de novo synthesis, then loss of umps-1 would block QA ability to rescue gonad delay in pnc-1 mutants. As expected, we found that QA did not rescue gonad delay in umps-1;pnc-1 double mutants, supporting UMPS-1’s role in de novo NAD+ synthesis. Moreover, loss of umps-1 decreased global NAD+ steady state levels compared to controls. Interestingly, loss of umps-1 blocked the incorporation of label from both Trp and QA into NAD+. UMPS-1 ability to use QA as a substrate for NAD+ biosynthesis supports an underground metabolic mechanism for NAD+ biosynthesis in a multicellular organism. Next, I would perform an enzymatic reaction expressing UMPS-1 and the substrate QA. I predict that this reaction would yield a NAD+ biosynthetic intermediate as the product if UMPS-1 were indeed using QA as a substrate for NAD+ biosynthesis. Furthermore, this suggests NAD+-associated underground 93 metabolism in higher organisms and supports an alternative approach to target NAD+ biosynthetic pathways for therapeutic potential. Therefore, there is a need to elucidate NAD+ homeostasis in mammalian models to maximize the efficiency of therapeutics boosting and inhibiting NAD+ biosynthetic pathways and metabolism.

NMRK-mediated synthesis in C. elegans

+ NR was recently discovered as a vitamin B3 precursor for NAD biosynthesis

(Bieganowski & Brenner, 2004). Although NMRK-mediated synthesis is highly conserved amongst organisms, there was not an annotated gene for NMRK-mediated synthesis in the C. elegans genome. Using BLAST, we identified T27A3.6 as the candidate enzyme for NMRK- mediated synthesis in C. elegans and renamed it NMRK-1. We discovered that we could reverse

NAD+-dependent phenotypes in pnc-1 mutants with both NR and NA supplementation. Loss of nmrk-1 decreased global NAD+ levels, supporting a role for NMRK-mediated synthesis in contributing to NAD+ biosynthesis. In addition to this, we uncovered a novel role for NMRK- mediated synthesis in C. elegans embryogenesis. Loss of nmrk-1 in combination with our UV- killed OP50 food source extends embryonic hatching for up to ten days. The ability of embryos to survive and hatch over that long period of time is remarkable. This supports the requirement of NAD+ biosynthetic capacity for reproduction, while illustrating diverse biological roles for each pathway. In order to identify key links connecting NMRK-mediated synthesis, UV-OP50 and embryonic hatching, I investigated embryonic hatching in the electron chain mutants, clk-1 and isp-1, which are known to have extended embryogenesis (Felkai et al., 1999; J. Feng et al.,

2001). Although embryonic hatching is slightly extended on the UV-OP50 food source, this does not compare to the ten days in nmrk-1 mutants. Furthermore, global metabolomics profiling (W.

Wang et al., 2015) suggested an increase in metabolites associated with oxidative stress on our 94

UV-killed OP50 food source. We were able to mimic the extended embryonic hatching phenotype in nmrk-1 mutants supplemented with paraquat. This supports the relationship between UV-killed OP50, oxidative stress and NMRK-mediated synthesis in C. elegans’ embryonic hatching. Next, we plan to investigate the components of embryonic hatching in order to identify the link with NMRK-mediated synthesis. Interestingly, NR is commonly found in cow’s milk, and is a beneficial dietary factor during development. Also, there have been therapeutic advances boosting NR to reverse various disease aliments, suggesting that NMRK- mediated synthesis is a promising drug target. Therefore, it is key to continue elucidating the biological impact of manipulating NMRK-mediated synthesis.

Homeostatic interactions amongst NAD+ biosynthetic pathways in C. elegans

Originally, we noticed an interesting trend amongst NMRK-mediated synthesis and de novo synthesis in loss of salvage NAD+ synthesis mutants. There was an increase expression of tdo-2 and nmrk-1 transcript levels in pnc-1 mutants. Not only did this trend support the contribution of each pathway to NAD+ biosynthetic capacity, but also it suggested a homeostatic response for the pathways in the absence of salvage NAD+ synthesis. I predicted that we could observe a reciprocal response in loss of kynu-1 and nmrk-1 mutants. As expected, I observed an increase of nmrk-1 and pnc-1 transcript levels in kynu-1 mutants, whereas, there was an increase in tdo-2 and pnc-1 in nmrk-1 mutants. This supports a compensatory network amongst NAD+ biosynthetic pathways to maintain NAD+ homeostasis. Therefore, this demonstrates a potential homeostatic mechanism in response to targeting NAD+ biosynthetic pathways for therapeutic use. For instance, inhibiting salvage NAD+ synthesis for anti-cancer therapeutics can result in increased NMRK-mediated synthesis and de novo synthesis. Whereas, targeting the kyneurine pathway in neurological disorders can result in increased NMRK-mediated synthesis and salvage 95

NAD+ synthesis. The kyneurine pathway is responsible for converting tryptophan into QA for

NAD+ biosynthesis. If de novo synthesis were manipulated, then what impact would this have on the kyneurine pathway and the neurological state? Furthermore, homeostatic interactions amongst NAD+ biosynthetic pathways should be taken into consideration when manipulating

NAD+ biosynthesis and metabolism for therapeutic benefits.

Compensatory network to maintain NAD+ homeostasis

The requirements for the cellular NAD+ pool are intensive. This metabolite is involved in cellular metabolic redox reactions. In addition to this responsibility, NAD+ serves as a substrate for a group of enzymes that regulate key biological processes. This suggests the need to maintain the NAD+ cellular pool for NAD+-dependent metabolic processes. Our loss of salvage NAD+ synthesis model suggests a compensatory mechanism that supplies carbon to the mitochondria when glycolysis is compromised. This supports homeostatic mechanisms are occurring to maintain cellular metabolism when NAD+ biosynthetic pathways are manipulated.

Targeting NAD+ biosynthetic pathways is an attractive route for anti-cancer and antibiotic therapeutics (Houtkooper & Auwerx, 2012; Murima et al., 2014; Pankiewicz et al.,

2015; A. A. Sauve, 2008; Srivastava, 2016). However, this work suggests that there are homeostatic responses amongst the pathways when another pathway is compromised. Therefore, therapeutics altering a certain biosynthetic pathway can lead to an additional response. Further investigation connecting NAD+ homeostatic requirements for cellular metabolism is required to elucidate the tissue and compartmental specific homeostatic responses when manipulating NAD+ biosynthetic pathways.

Our model also revealed that boosting NAD+ precursors could reverse NAD+-dependent phenotypes in pnc-1 mutants. NAD+ decline is associated with aging and age-related metabolic 96 disorders (Gomes et al., 2013; Mills et al., 2016; Mouchiroud et al., 2013). Boosting NAD+ in various models can reverse these age-associated defects (Gomes et al., 2013; Houtkooper &

Auwerx, 2012; Mills et al., 2016; Mouchiroud et al., 2013; Srivastava, 2016). This suggests maintaining NAD+ homeostasis can promote aging healthier. Therapeutics designed to boost

NAD+ biosynthesis has the potential of declining metabolic aliments due to aging induced NAD+ decline.

NAD+ precursors and PARP inhibitors are neuroprotective (Fang et al., 2014; Klaidman et al., 2003; Qin et al., 2006; L. Wang et al., 2014). PARPs are considered to consume a large amount of NAD+ for DNA damage response (Chiarugi, 2012; Zhou et al., 2015). Therefore, it’s no surprise that over-activation of PARPs are observed in neurological disease states. Boosting

NAD+ biosynthesis, via precursor supplementation, can protect against neuronal axon injury,

Alzheimer’s disease aliments and hearing loss (Brown et al., 2014; Gerdts et al., 2015; Gong et al., 2013; Qin et al., 2006; Sasaki et al., 2006; L. Wang et al., 2014). This supports the requirement of NAD+ homeostasis and maintaining NAD+ cellular pools for normal neurological progression and protection against neurological aliments and decline. Notably, the kyneurine pathway is associated with neurological processes and required for converting tryptophan into

NAD+ via de novo synthesis (Breda et al., 2016; Erhardt et al., 2016; Lim et al., 2015; Majewski et al., 2016; Schwarcz & Pellicciari, 2002; Vamos et al., 2009). This suggests manipulating

NAD+ biosynthetic pathways could result in adverse results. Therefore, future therapeutics targeting NAD+ biosynthetic pathways should consider the metabolic homeostatic interactions due to manipulation.

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Unique and diverse biological functions for NAD+ biosynthetic pathways involved in C. elegans reproduction

Our lab previously uncovered a novel role for salvage NAD+ synthesis in C. elegans’s gonad development (Huang & Hanna-Rose, 2006; T L Vrablik et al., 2009; Tracy L Vrablik et al., 2011). We were able to further link the impairment of reproductive development to dysregulated glycolysis in pnc-1 mutants (W. Wang et al., 2015). There are also diverse phenotypes associated with NAM accumulation in pnc-1 mutants (Upadhyay et al., 2016; T L

Vrablik et al., 2009). The requirement of salvage synthesis for normal timing of gonad development suggests separable biological roles for NMRK-mediated and de novo synthesis in

C. elegans. I identified a link between de novo NAD+ synthesis and fecundity in C. elegans. Loss of kynu-1 decreased progeny production compared to controls. QA supplementation reversed the progeny production decline in kynu-1 mutants. This suggests a dynamic biological role for either the kyneurine pathway or de novo NAD+ synthesis in regulating fecundity in C. elegans.

Surprisingly, I also discovered a key unique role for NMRK-mediated synthesis in embryonic hatching. Embryos survive and hatch over time for up to ten days in our nmrk-1 mutants fed

UV-killed OP50. We were able to further mimic the affect of UV-killed OP50 on hatching in nmrk-1 mutants exposed to oxidative stress. This connects NMRK-mediated synthesis, oxidative stress and embryonic hatching in C. elegans. The novel role of NMRK-mediated synthesis in embryonic hatching supports the role of NMRK-mediated NAD+ biosynthesis in C. elegans’ reproduction. Interestingly, extended embryonic hatching was not observed in de novo NAD+ synthesis and salvage NAD+ synthesis mutants. I predict that the distinct biological roles for

NAD+ biosynthetic pathways are due to specific tissue and compartmental needs for NAD+ biosynthetic capacity. Elucidating the tissue and compartmental-specific requirement for NAD+ 98 biosynthesis is critical for uncovering the mechanisms behind NAD+ homeostasis and distinct biological roles.

It’s interesting to note that although each NAD+ biosynthetic pathway has diverse and unique biological roles, the roles are still associated with reproduction. This phenomenon could be accounted for by one or two explanations. First, the energy requirement during reproduction competes with metabolic energy requirements. Manipulating each NAD+ biosynthetic pathway results in a metabolic shift that maintains functions of the mitochondria. This suggests that an aspect of reproduction lack in function when a NAD+ biosynthetic pathway is compromised.

Secondly, this suggests that NAD+ biosynthesis is absolutely required for normal progression of reproduction. I predict that both explanations illustrate what’s occurring. Elucidating the energy requirements required for reproduction when each NAD+ biosynthetic pathway is compromised will shed light on underlying this mechanism. This supports the novel and diverse biological roles for NAD+ biosynthetic pathways.

Underground metabolism: Alternative routes to synthesize NAD+

It’s understood that the biochemical mechanisms of NAD+ biosynthesis and metabolism are well defined and studied, first identified in 1906; however, a newly found vitamin B3 precursor, NR, was recently discovered (Bieganowski & Brenner, 2004). Since its discovery, NR and NMRK-mediated synthesis has shown to have numerous biological roles and the capacity to protect against many disease states (Cantó et al., 2012; Gariani et al., 2016; Lee et al., 2015;

Ratajczak et al., 2016; Trammell et al., 2016). Identifying NR as an additional vitamin B3 precursor greatly impacted both the NAD+ field and therapeutic approaches (Houtkooper &

Auwerx, 2012; Mills et al., 2016; Mouchiroud et al., 2013; A. A. Sauve, 2008; Srivastava, 2016;

Y. Yang & Sauve, 2016). This supports the need to further elucidate the mechanisms behind 99

NAD+ homeostasis. Also, this suggests there could be alternative enzymes using NAD+ intermediates as substrates. For example, we discovered that UMPS-1 could substitute for

QPRTase for NAD+ biosynthesis in C. elegans. This supports the role of underground metabolism illustrating the metabolic plasticity of UMPS-1 and de novo NAD+ biosynthesis

(D’Ari & Casadesus, 1998).

100

References:

Altschul, R., Hoffer, A., & Stephen, J. D. (1955). Influence of Nicotinic Acid on Serum Cholesterol in Man. Arch. Biochem. Biophys., 54(2), 558–559. https://doi.org/10.1016/0003-9861(55)90070-9 Antebi, A., Yeh, W. H., Tait, D., Hedgecock, E. M., & Riddle, D. L. (2000). daf-12 encodes a nuclear receptor that regulates the dauer diapause and developmental age in C. elegans. Genes and Development, 14(12), 1512–1527. https://doi.org/10.1101/gad.14.12.1512 Antebi, a, Culotti, J. G., & Hedgecock, E. M. (1998). daf-12 regulates developmental age and the dauer alternative in Caenorhabditis elegans. Development (Cambridge, England), 125(7), 1191–1205. Bai, P., Canto, C., Oudart, H., Brunyánszki, A., Cen, Y., Thomas, C., … Auwerx, J. (2011). PARP-1 inhibition increases mitochondrial metabolism through SIRT1 activation. April, 6(134), 461–468. https://doi.org/10.1016/j.cmet.2011.03.004 Ball, H. J., Yuasa, H. J., Austin, C. J. D., Weiser, S., & Hunt, N. H. (2009). Indoleamine 2,3- dioxygenase-2; a new enzyme in the kynurenine pathway. International Journal of Biochemistry and Cell Biology, 41(3), 467–471. https://doi.org/10.1016/j.biocel.2008.01.005 Baur, J. A., Pearson, K. J., Price, N. L., Jamieson, H. A., Lerin, C., Kalra, A., … Sinclair, D. A. (2006). Resveratrol improves health and survival of mice on a high-calorie diet. Nature, 444(7117), 337–342. https://doi.org/10.1038/nature05354 Belenky, P., Bogan, K. L., & Brenner, C. (2007). NAD+ metabolism in health and disease. Trends in Biochemical Sciences, 32(1), 12–19. https://doi.org/10.1016/j.tibs.2006.11.006 Benavente, C. A., Schnell, S. A., & Jacobson, E. L. (2012). Effects of Niacin restriction on sirtuin and PARP responses to photodamage in human skin. PLoS ONE, 7(7). https://doi.org/10.1371/journal.pone.0042276 Benyo, Z., Gille, A., Kero, J., Csiky, M., Suchankova, M. C., Nusing, R. M., … Offermanns, S. (n.d.). GRP109A (PUMA-G/HM74A) mediates nicotinic acid-induced flushing. Bieganowski, P., & Brenner, C. (2004). Discoveries of nicotinamide riboside as a nutrient and conserved NRK genes establish a Preiss-Handler independent route to NAD+ in fungi and humans. Cell, 117(4), 495–502. Retrieved from http://www.ncbi.nlm.nih.gov/pubmed/15137942 Birjmohun, R. S., Hutten, B. A., Kastelein, J. J. P., & Stroes, E. S. G. (2005). Efficacy and safety of high-density lipoprotein cholesterol-increasing compounds: A meta-analysis of randomized controlled trials. Journal of the American College of Cardiology, 45(2), 185– 197. https://doi.org/10.1016/j.jacc.2004.10.031 Bogan, K. L., & Brenner, C. (2008a). Nicotinic acid, nicotinamide, and nicotinamide riboside: a molecular evaluation of NAD+ precursor vitamins in human nutrition. Annual Review of Nutrition, 28, 115–30. https://doi.org/10.1146/annurev.nutr.28.061807.155443 Bogan, K. L., & Brenner, C. (2008b). Nicotinic acid, nicotinamide, and nicotinamide riboside: a molecular evaluation of NAD+ precursor vitamins in human nutrition. Annual Review of Nutrition, 28, 115–130. https://doi.org/10.1146/annurev.nutr.28.061807.155443 Bonkowski, M. S., & Sinclair, D. A. (2016). Slowing ageing by design: the rise of NAD+ and sirtuin-activating compounds. Nature Reviews Molecular Cell Biology, 230(2001), 2–3. https://doi.org/10.1038/nrm.2016.93 Braidy, N., Guillemin, G. J., Mansour, H., Chan-Ling, T., Poljak, A., & Grant, R. (2011). Age 101

related changes in NAD+ metabolism oxidative stress and sirt1 activity in wistar rats. PLoS ONE, 6(4), 1–18. https://doi.org/10.1371/journal.pone.0019194 Breda, C., Sathyasaikumar, K. V., Sograte Idrissi, S., Notarangelo, F. M., Estranero, J. G., Moore, G. G. L., … Giorgini, F. (2016). Tryptophan-2,3-dioxygenase (TDO) inhibition ameliorates neurodegeneration by modulation of kynurenine pathway metabolites. Proceedings of the National Academy of Sciences of the United States of America, 113(14). https://doi.org/10.1073/pnas.1604453113 Brenner, C. (2014). Boosting NAD to spare hearing. Cell Metabolism, 20(6), 926–927. https://doi.org/10.1016/j.cmet.2014.11.015 Brenner, S. (1974). Caenorhabdztzs elegans. Methods, 77(1), 71–94. https://doi.org/10.1111/j.1749-6632.1999.tb07894.x Brown, K. D., Maqsood, S., Huang, J. Y., Pan, Y., Harkcom, W., Li, W., … Jaffrey, S. R. (2014). Activation of SIRT3 by the NAD+ precursor nicotinamide riboside protects from noise-induced hearing loss. Cell Metabolism, 20(6), 1059–1068. https://doi.org/10.1016/j.cmet.2014.11.003 Cant??, C., Menzies, K. J., & Auwerx, J. (2015). NAD+ Metabolism and the Control of Energy Homeostasis: A Balancing Act between Mitochondria and the Nucleus. Cell Metabolism, 22(1), 31–53. https://doi.org/10.1016/j.cmet.2015.05.023 Canto, C., & Auwerx, J. (2012). Targeting sirtuin 1 to improve metabolism: all you need is NAD(+)? Pharmacol Rev, 64(1), 166–187. https://doi.org/10.1124/pr.110.003905 Cantó, C., Houtkooper, R. H., Pirinen, E., Youn, D. Y., Oosterveer, M. H., Cen, Y., … Auwerx, J. (2012). The NAD(+) Precursor Nicotinamide Riboside Enhances Oxidative Metabolism and Protects against High-Fat Diet-Induced Obesity. Cell Metabolism, 15(6), 838–47. https://doi.org/10.1016/j.cmet.2012.04.022 Cerutti, R., Pirinen, E., Lamperti, C., Marchet, S., Sauve, A. A., Li, W., … Zeviani, M. (2014). NAD+-dependent activation of Sirt1 corrects the phenotype in a mouse model of mitochondrial disease. Cell Metabolism, 19(6), 1042–1049. https://doi.org/10.1016/j.cmet.2014.04.001 Chen, Y., & Guillemin, G. J. (2009). Kynurenine pathway metabolites in humans: Disease and healthy states. International Journal of Tryptophan Research, 2(1), 1–19. https://doi.org/10.4137/ijtr.s2097 Chi, Y., & Sauve, A. a. (2013). Nicotinamide riboside, a trace nutrient in foods, is a vitamin B3 with effects on energy metabolism and neuroprotection. Current Opinion in Clinical Nutrition and Metabolic Care, 16(6). https://doi.org/10.1097/MCO.0b013e32836510c0 Chiarugi, A. (2012). A snapshot of chemoresistance to PARP inhibitors. Trends in Pharmacological Sciences, 33(1), 42–48. https://doi.org/10.1016/j.tips.2011.10.001 Chiarugi, A., Dolle, C., Felici, R., Ziegler, M., Dölle, C., Felici, R., … Ziegler, M. (2012). The NAD metabolome--a key determinant of cancer cell biology. Nature Reviews. Cancer, 12(11), 741–52. https://doi.org/10.1038/nrc3340 D’Ari, R., & Casades??s, J. (1998). Underground metabolism. BioEssays, 20(2), 181–186. https://doi.org/10.1002/(SICI)1521-1878(199802)20:2<181::AID-BIES10>3.0.CO;2-0 de Figueiredo, L. F., Gossmann, T. I., Ziegler, M., & Schuster, S. (2011a). Pathway analysis of NAD+ metabolism. The Biochemical Journal, 439(2), 341–8. https://doi.org/10.1042/BJ20110320 de Figueiredo, L. F., Gossmann, T. I., Ziegler, M., & Schuster, S. (2011b). Pathway analysis of NAD+ metabolism. The Biochemical Journal, 439(2), 341–8. 102

https://doi.org/10.1042/BJ20110320 Erhardt, S., Schwieler, L., Imbeault, S., & Engberg, G. (2016). The kynurenine pathway in schizophrenia and bipolar disorder. Neuropharmacology. https://doi.org/10.1016/j.neuropharm.2016.05.020 Falk, M. J., Rao, M., Ostrovsky, J., Daikhin, E., Nissim, I., & Yudkoff, M. (2011). Stable isotopic profiling of intermediary metabolic flux in developing and adult stage Caenorhabditis elegans. Journal of Visualized Experiments : JoVE, (48), 1–7. https://doi.org/10.3791/2288 Fang, E. F., Scheibye-Knudsen, M., Brace, L. E., Kassahun, H., Sengupta, T., Nilsen, H., … Bohr, V. A. (2014). Defective mitophagy in XPA via PARP-1 hyperactivation and NAD +/SIRT1 reduction. Cell, 157(4), 882–896. https://doi.org/10.1016/j.cell.2014.03.026 Fatokun, A. A., Hunt, N. H., & Ball, H. J. (2013). Indoleamine 2,3-dioxygenase 2 (IDO2) and the kynurenine pathway: Characteristics and potential roles in health and disease. Amino Acids. https://doi.org/10.1007/s00726-013-1602-1 Felkai, S., Ewbank, J. J., Lemieux, J., Labbé, J. C., Brown, G. G., & Hekimi, S. (1999). CLK-1 controls respiration, behavior and aging in the nematode Caenorhabditis elegans. The EMBO Journal, 18(7), 1783–1792. https://doi.org/10.1093/emboj/18.7.1783 Feng, J., Bussière, F., & Hekimi, S. (2001). Mitochondrial electron transport is a key determinant of life span in Caenorhabditis elegans. Developmental Cell, 1(5), 633–644. https://doi.org/10.1016/S1534-5807(01)00071-5 Feng, Y., Paul, I. A., & LeBlanc, M. H. (2006). Nicotinamide reduces hypoxic ischemic brain injury in the newborn rat. Brain Research Bulletin, 69(2), 117–122. https://doi.org/10.1016/j.brainresbull.2005.11.011 French, J. B., Cen, Y., Vrablik, T. L., Xu, P., Allen, E., Hanna-Rose, W., & Sauve, A. a. (2010). Characterization of nicotinamidases: steady state kinetic parameters, classwide inhibition by nicotinaldehydes, and catalytic mechanism. Biochemistry, 49(49), 10421–39. https://doi.org/10.1021/bi1012518 Gariani, K., Menzies, K. J., Ryu, D., Wegner, C. J., Wang, X., Ropelle, E. R., … Auwerx, J. (2016). Eliciting the mitochondrial unfolded protein response by nicotinamide adenine dinucleotide repletion reverses fatty liver disease in mice. Hepatology, 63(4), 1190–1204. https://doi.org/10.1002/hep.28245 Gensler, H. L., Williams, T., Huang, A. C., & Jacobson, E. L. (1999). Oral niacin prevents photocarcinogenesis and photoimmunosuppression in mice. Nutr Cancer, 34(1), 36–41. https://doi.org/10.1207/S15327914NC340105 Gerdts, J., Brace, E. J., Sasaki, Y., Diantonio, A., & Milbrandt, J. (2015). SARM1 activation triggers axon degeneration locally via NAD+ destruction. Gerisch, B., Weitzel, C., Kober-Eisermann, C., Rottiers, V., & Antebi, A. (2001). A Hormonal Signaling Pathway Influencing C. elegans Metabolism, Reproductive Development, and Life Span. Developmental Cell, 1(6), 841–851. https://doi.org/10.1016/S1534- 5807(01)00085-5 Goldstein, L. E., Leopold, M. C., Huang, X., Atwood, C. S., Saunders, A. J., Hartshorn, M., … Bush, A. I. (2000). 3-Hydroxykynurenine and 3-Hydroxyanthranilic Acid Generate Hydrogen Peroxide and Promote ??-Crystallin Cross-Linking By Metal Ion Reduction. Biochemistry, 39(24), 7266–7275. https://doi.org/10.1021/bi992997s Gomes, A. P., Price, N. L., Ling, A. J. Y., Moslehi, J. J., Montgomery, M. K., Rajman, L., … Sinclair, D. A. (2013). Declining NAD+ induces a pseudohypoxic state disrupting nuclear- 103

mitochondrial communication during aging. Cell, 155(7). https://doi.org/10.1016/j.cell.2013.11.037 Gong, B., Pan, Y., Vempati, P., Zhao, W., Knable, L., Ho, L., … Pasinetti, G. M. (2013). Nicotinamide riboside restores cognition through an upregulation of proliferator-activated receptor-γ coactivator 1α regulated β-secretase 1 degradation and mitochondrial gene expression in Alzheimer’s mouse models. Neurobiology of Aging, 34(6), 1581–1588. https://doi.org/10.1016/j.neurobiolaging.2012.12.005 Gossmann, T. I., Ziegler, M., Puntervoll, P., De Figueiredo, L. F., Schuster, S., & Heiland, I. (2012a). NAD(+) biosynthesis and salvage - a phylogenetic perspective. The FEBS Journal, 279(18), 1–9. https://doi.org/10.1111/j.1742-4658.2012.08559.x Gossmann, T. I., Ziegler, M., Puntervoll, P., De Figueiredo, L. F., Schuster, S., & Heiland, I. (2012b). NAD+ biosynthesis and salvage - A phylogenetic perspective. FEBS Journal, 279(18), 3355–3363. https://doi.org/10.1111/j.1742-4658.2012.08559.x Gu, W., Wang, T., Maltais, F., Ledford, B., Kennedy, J., Wei, Y., … Charifson, P. S. (2012). Design, synthesis and biological evaluation of potent NAD+-dependent DNA ligase inhibitors as potential antibacterial agents. Part I: aminoalkoxypyrimidine carboxamides. Bioorg. Med. Chem. Lett., 22(11), 3693–3698. https://doi.org/10.1016/j.bmcl.2012.04.037 Guse, A. H. (2005). Second messenger function and the structure-activity relationship of cyclic adenosine diphosphoribose (cADPR). FEBS Journal, 272(18), 4590–4597. https://doi.org/10.1111/j.1742-4658.2005.04863.x Gwirtz, J. A., & Garcia-Casal, M. N. (n.d.). Issue: Technical Considerations for Maize Flour and Corn Meal Fortification in Public Health Processing maize flour and corn meal food products. Ann. N.Y. Acad. Sci. https://doi.org/10.1111/nyas.12299 Haigis, M. C., Sinclair, D. A., & Edu, M. H. (n.d.). Mammalian Sirtuins: Biological Insights and Disease Relevance. https://doi.org/10.1146/annurev.pathol.4.110807.092250 Harris, F., & Pierpoint, L. (2012). PhotodynamicTherapy Based on 5-Aminolevulinic Acid and Its Use as an Antimicrobial Agent. Medicinal Research Reviews, 29(6), 1292–1327. https://doi.org/10.1002/med Hasmann, M., & Schemainda, I. (2003). FK866, a Highly Specific Noncompetitive Inhibitor of Nicotinamide Phosphoribosyltransferase, Represents a Novel Mechanism for Induction of Tumor Cell Apoptosis. Cancer Research, 63(21), 7436–7442. Hegyi, J., Schwartz, R. A., & Hegyi, V. (2004). Pellagra : Dermatitis , dementia , and diarrhea, 1–5. Herranz, D., Muñoz-Martin, M., Cañamero, M., Mulero, F., Martinez-Pastor, B., Fernandez- Capetillo, O., & Serrano, M. (2010). Sirt1 improves healthy ageing and protects from metabolic syndrome-associated cancer. Nature Communications, 1, 3. https://doi.org/10.1038/ncomms1001 Hoogewijs, D., Houthoofd, K., Matthijssens, F., Vandesompele, J., & Vanfleteren, J. R. (2008). Selection and validation of a set of reliable reference genes for quantitative sod gene expression analysis in C. elegans. BMC Molecular Biology, 9, 9. https://doi.org/10.1186/1471-2199-9-9 Houtkooper, R. H., & Auwerx, J. (2012). Exploring the therapeutic space around NAD+. Journal of Cell Biology, 199(2), 205–209. https://doi.org/10.1083/jcb.201207019 Houtkooper, R. H., Cantó, C., Wanders, R. J., & Auwerx, J. (2010). The secret life of NAD+: an old metabolite controlling new metabolic signaling pathways. Endocrine Reviews, 31(2), 194–223. https://doi.org/10.1210/er.2009-0026 104

Huang, L., & Hanna-Rose, W. (2006). EGF signaling overcomes a uterine cell death associated with temporal mis-coordination of organogenesis within the C. elegans egg-laying apparatus. Developmental Biology, 300(2), 599–611. https://doi.org/10.1016/j.ydbio.2006.08.024 Ieraci, A., & Herrera, D. G. (2006). Nicotinamide protects against ethanol-induced apoptotic neurodegeneration in the developing mouse brain. PLoS Medicine, 3(4), 547–557. https://doi.org/10.1371/journal.pmed.0030101 Jack Preiss, P. H. (1957). Enzymatic Synthesis of. Biological Chemistry. Jhamandas, K., Boegman, R. J., Beninger, R. J., & Bialik, M. (1990). Quinolinate-induced cortical cholinergic damage: modulation by tryptophan metabolites. Brain Research, 529(1– 2), 185–191. https://doi.org/10.1016/0006-8993(90)90826-W Khan, N. A., Auranen, M., Paetau, I., Pirinen, E., Euro, L., Forsström, S., … DiDonato, S. (2014). Effective treatment of mitochondrial myopathy by nicotinamide riboside, a vitamin B3. EMBO Molecular Medicine, 6(6), n/a-n/a. https://doi.org/10.1002/emmm.201403943 Kim, J.-H., O’Brien, K. M., Sharma, R., Boshoff, H. I. M., Rehren, G., Chakraborty, S., … Schnappinger, D. (2013). A genetic strategy to identify targets for the development of drugs that prevent bacterial persistence. Pnas, 110(47), 19095–19100. https://doi.org/10.1073/pnas.1315860110 Klaidman, L., Morales, M., Kem, S., Yang, J., Chang, M. L., & Adams, J. D. (2003). Nicotinamide offers multiple protective mechanisms in stroke as a precursor for NAD+, as a PARP inhibitor and by partial restoration of mitochondrial function. Pharmacology, 69(3), 150–157. https://doi.org/10.1159/000072668 Koppenol, W. H., Bounds, P. L., & Dang, C. V. (2011). Otto Warburg’s contributions to current concepts of cancer metabolism. Nature Reviews. Cancer, 11(5), 325–37. https://doi.org/10.1038/nrc3038 Lagouge, M., Argmann, C., Gerhart-Hines, Z., Meziane, H., Lerin, C., Daussin, F., … Auwerx, J. (2006). Resveratrol Improves Mitochondrial Function and Protects against Metabolic Disease by Activating SIRT1 and PGC-1?? Cell, 127(6), 1109–1122. https://doi.org/10.1016/j.cell.2006.11.013 Lee, H. J., Hong, Y.-S., Jun, W., & Yang, S. J. (2015). Nicotinamide Riboside Ameliorates Hepatic Metaflammation by Modulating NLRP3 Inflammasome in a Rodent Model of Type 2 Diabetes. Journal of Medicinal Food, 0(0), 150514130725000. https://doi.org/10.1089/jmf.2015.3439 Lim, C. K., Fern??ndez-Gomez, F. J., Braidy, N., Estrada, C., Costa, C., Costa, S., … Guillemin, G. J. (2015). Involvement of the kynurenine pathway in the pathogenesis of Parkinson’s disease. Progress in Neurobiology, (2015). https://doi.org/10.1016/j.pneurobio.2015.12.009 Lu, W., Clasquin, M. F., Melamud, E., Amador-Noguez, D., Caudy, A. A., & Rabinowitz, J. D. (2010). Metabolomic analysis via reversed-phase ion-pairing liquid chromatography coupled to a stand alone orbitrap mass spectrometer. Analytical Chemistry, 82, 3212–3221. https://doi.org/10.1021/ac902837x Magni, G., Amici, A., Emanuelli, M., Orsomando, G., Raffaelli, N., & Ruggieri, S. (2004). Enzymology of NAD+ homeostasis in man. Cellular and Molecular Life Sciences, 61(1), 19–34. https://doi.org/10.1007/s00018-003-3161-1 Majewski, M., Kozlowska, A., Thoene, M., Lepiarczyk, E., & Grzegorzewski, W. J. (2016). Overview of the role of vitamins and minerals on the kynurenine pathway in health and disease. Journal of Physiology and Pharmacology, 67(1), 3–20. 105

Massudi, H., Grant, R., Braidy, N., Guest, J., Farnsworth, B., & Guillemin, G. J. (2012). Age- associated changes in oxidative stress and NAD+ metabolism in human tissue. PLoS ONE, 7(7), 1–9. https://doi.org/10.1371/journal.pone.0042357 Mills, K. F., Yoshida, S., Stein, L. R., Grozio, A., Kubota, S., Sasaki, Y., … Imai, S. (2016). Long-Term Administration of Nicotinamide Mononucleotide Mitigates Age-Associated Physiological Decline in Mice. Cell Metabolism, 24(6), 1–12. https://doi.org/10.1016/j.cmet.2016.09.013 Mouchiroud, L., Houtkooper, R. H., & Auwerx, J. (2013). NAD+ metabolism: a therapeutic target for age-related metabolic disease. Critical Reviews in Biochemistry and Molecular Biology, 48(4), 397–408. https://doi.org/10.3109/10409238.2013.789479 Murima, P., McKinney, J. D., & Pethe, K. (2014). Targeting bacterial central metabolism for drug development. Chemistry and Biology, 21(11), 1423–1432. https://doi.org/10.1016/j.chembiol.2014.08.020 O’Farrell, K., & Harkin, A. (2017). Stress-related regulation of the kynurenine pathway: Relevance to neuropsychiatric and degenerative disorders. Neuropharmacology. https://doi.org/10.1016/j.neuropharm.2015.12.004 Ohashi, K., Kawai, S., & Murata, K. (2013). Secretion of quinolinic acid, an intermediate in the kynurenine pathway, for utilization in NAD+ biosynthesis in the yeast Saccharomyces cerevisiae. Eukaryotic Cell, 12(5), 648–653. https://doi.org/10.1128/EC.00339-12 Oxenkrug, G. F. (2010). Tryptophan kynurenine metabolism as a common mediator of genetic and environmental impacts in major depressive disorder: the serotonin hypothesis revisited 40 years later. The Israel Journal of Psychiatry and Related Sciences, 47(1), 56–63. https://doi.org/10.1016/j.neulet.2010.11.003.Melatonin Pankiewicz, K. W., Petrelli, R., Singh, R., & Felczak, K. (2015). Nicotinamide Adenine Dinucleotide Based Therapeutics, Update. Current Medicinal Chemistry, 22(34), 3991– 4028. https://doi.org/10.2174/0929867322666150821100720 Perkins, M. N., & Stone, T. W. (1982). An iontophoretic investigation of the actions of convulsant kynurenines and their interaction with the endogenous excitant quinolinic acid. Brain Research, 247(1), 184–187. https://doi.org/10.1016/0006-8993(82)91048-4 Pirinen, E., Canto, C., Jo, Y.-S., Morato, L., Zhang, H., Menzies, K., … Auwerx, J. (2014). Pharmacological Inhibition of Poly(ADP-Ribose) Polymerases Improves Fitness and Mitochondrial Function in Skeletal Muscle. Cell Metab, 19(6), 1034–1041. https://doi.org/10.1016/j.cmet.2014.04.002 Pittelli, M., Felici, R., Pitozzi, V., Giovannelli, L., Bigagli, E., Cialdai, F., … Chiarugi, A. (n.d.). Pharmacological Effects of Exogenous NAD on Mitochondrial Bioenergetics, DNA Repair, and Apoptosis. https://doi.org/10.1124/mol.111.073916 Qin, W., Yang, T., Ho, L., Zhao, Z., Wang, J., Chen, L., … Pasinetti, G. M. (2006). Neuronal SIRT1 activation as a novel mechanism underlying the prevention of alzheimer disease amyloid neuropathology by calorie restriction. Journal of Biological Chemistry, 281(31), 21745–21754. https://doi.org/10.1074/jbc.M602909200 Rajakumar, K. (2000). Pellagra in the United States: a historical perspective. Southern Medical Journal, 93(3), 272–7. Retrieved from http://www.ncbi.nlm.nih.gov/pubmed/10728513 Ratajczak, J., Joffraud, M., Trammell, S. a. J., Ras, R., Canela, N., Boutant, M., … Cantó, C. (2016). NRK1 controls nicotinamide mononucleotide and nicotinamide riboside metabolism in mammalian cells. Nature Communications, 7, 13103. https://doi.org/10.1038/ncomms13103 106

Rittershaus, E. S. C., Baek, S. H., & Sassetti, C. M. (2013). The normalcy of dormancy: Common themes in microbial quiescence. Cell Host and Microbe, 13(6), 643–651. https://doi.org/10.1016/j.chom.2013.05.012 Rongvaux, A., Andris, F., Van Gool, F., & Leo, O. (2003). Reconstructing eukaryotic NAD metabolism. BioEssays, 25(7), 683–690. https://doi.org/10.1002/bies.10297 Rowent, J. W. (1951). The phosphorolysis, 210. Sadanaga-Akiyoshi, F., Yao, H., Tanuma, S. ichi, Nakahara, T., Hong, J. S., Ibayashi, S., … Fujishima, M. (2003). Nicotinamide attenuates focal ischemic brain injury in rats: With special reference to changes in nicotinamide and NAD+ levels in ischemic core and penumbra. Neurochemical Research, 28(8), 1227–1234. https://doi.org/10.1023/A:1024236614015 Santidrian, A. F., Matsuno-yagi, A., Ritland, M., Seo, B. B., Leboeuf, S. E., Gay, L. J., … Felding-habermann, B. (2013). Mitochondrial complex I activity and NAD + / NADH balance regulate breast cancer progression. The Journal of Clinical Investigation, 123(3), 1068–1081. https://doi.org/10.1172/JCI64264DS1 Sas, K., Robotka, H., Toldi, J., & Vécsei, L. (2007a). Mitochondria, metabolic disturbances, oxidative stress and the kynurenine system, with focus on neurodegenerative disorders. Journal of the Neurological Sciences, 257(1–2), 221–239. https://doi.org/10.1016/j.jns.2007.01.033 Sas, K., Robotka, H., Toldi, J., & Vécsei, L. (2007b). Mitochondria, metabolic disturbances, oxidative stress and the kynurenine system, with focus on neurodegenerative disorders. Journal of the Neurological Sciences, 257(1–2), 221–39. https://doi.org/10.1016/j.jns.2007.01.033 Sasaki, Y., Araki, T., & Milbrandt, J. (2006). Stimulation of nicotinamide adenine dinucleotide biosynthetic pathways delays axonal degeneration after axotomy. The Journal of Neuroscience : The Official Journal of the Society for Neuroscience, 26(33), 8484–91. https://doi.org/10.1523/JNEUROSCI.2320-06.2006 Sauve, A. a. (2008). NAD + and Vitamin B 3 : From Metabolism to Therapies. The Journal of Pharmacology and Experimental Therapeutics., 324(3), 883–893. https://doi.org/10.1124/jpet.107.120758.energy Sauve, A. A. (2008). NAD ϩ and Vitamin B 3 : From Metabolism to Therapies, 324(3), 883–893. https://doi.org/10.1124/jpet.107.120758.energy Scheibye-Knudsen, M., Fang, E. F., Croteau, D. L., & Bohr, V. A. (2014). Contribution of defective mitophagy to the neurodegeneration in DNA repair-deficient disorders. Autophagy, 10(8), 1468–1469. https://doi.org/10.4161/auto.29321 Schrier Vergano, S., Rao, M., McCormack, S., Ostrovsky, J., Clarke, C., Preston, J., … Falk, M. J. (2014). In vivo metabolic flux profiling with stable isotopes discriminates sites and quantifies effects of mitochondrial dysfunction in C. elegans. Molecular Genetics and Metabolism, 111(3), 331–341. https://doi.org/10.1016/j.ymgme.2013.12.011 Schwarcz, R., & Pellicciari, R. (2002). Manipulation of brain kynurenines: glial targets, neuronal effects, and clinical opportunities. The Journal of Pharmacology and Experimental Therapeutics, 303(1), 1–10. https://doi.org/10.1124/jpet.102.034439 Shuman, S. (2009). DNA ligases: Progress and prospects. Journal of Biological Chemistry, 284(26), 17365–17369. https://doi.org/10.1074/jbc.R900017200 Society, T. R., Society, R., Papers, C., & Character, B. (2011). The Alcoholic Ferment of Yeast- Juice . Part V . -The Function of Phosphates in Alcoholic Fermentation Author ( s ): Arthur 107

Harden and William John Young Source : Proceedings of the Royal Society of London . Series B , Containing Papers of a Published by : Society, 82(556), 321–330. Srivastava, S. (2016). Emerging therapeutic roles for NAD+ metabolism in mitochondrial and age-related disorders. Clinical and Translational Medicine, 5(1), 25. https://doi.org/10.1186/s40169-016-0104-7 Stokes, S. S., Gowravaram, M., Huynh, H., Lu, M., Mullen, G. B., Chen, B., … Mills, S. D. (2012). Discovery of bacterial NAD +-dependent DNA ligase inhibitors: Improvements in clearance of adenosine series. Bioorganic and Medicinal Chemistry Letters, 22(1), 85–89. https://doi.org/10.1016/j.bmcl.2011.11.071 Stone, T. W., & Perkins, M. N. (1981). Quinolinic acid: a potent endogenous excitant at amino acid receptors in CNS. European Journal of Pharmacology, 72(4), 411–2. Sydenstricker, V. P. (n.d.). The History of Pellagra, Its Recognition as a Disorder of Nutrition and Its Conquest. Tateishi, K., Wakimoto, H., Iafrate, A. J., Tanaka, S., Loebel, F., Lelic, N., … Cahill, D. P. (2015). Extreme Vulnerability of IDH1 Mutant Cancers to NAD+ Depletion. Cancer Cell, 28(6), 773–784. https://doi.org/10.1016/j.ccell.2015.11.006 Timson, D. J., Singleton, M. R., & Wigley, D. B. (2000). DNA ligases in the repair and replication of DNA. Mutation Research - DNA Repair, 460(3–4), 301–318. https://doi.org/10.1016/S0921-8777(00)00033-1 Trammell, S. A. J., Schmidt, M. S., Weidemann, B. J., Redpath, P., Jaksch, F., Dellinger, R. W., … Lau, D. T. (2016). Nicotinamide riboside is uniquely and orally bioavailable in mice and humans. Nature Communications, 7, 12948. https://doi.org/10.1038/ncomms12948 Tummala, K. S., Gomes, A. L., Yilmaz, M., Graña, O., Bakiri, L., Ruppen, I., … Djouder, N. (2014). Inhibition of De Novo NAD+ Synthesis by Oncogenic URI Causes Liver Tumorigenesis through DNA Damage. Cancer Cell, 26(6), 826–839. https://doi.org/10.1016/j.ccell.2014.10.002 Turunc Bayrakdar, E., Uyanikgil, Y., Kanit, L., Koylu, E., & Yalcin, a. (2014). Nicotinamide treatment reduces the levels of oxidative stress, apoptosis, and PARP-1 activity in Aβ(1- 42)-induced rat model of Alzheimer’s disease. Free Radical Research, 48(2), 146–58. https://doi.org/10.3109/10715762.2013.857018 Upadhyay, A., Pisupati, A., Jegla, T., Crook, M., Mickolajczyk, K. J., Shorey, M., … Hanna- Rose, W. (2016). Vitamin B3 / Nicotinamide is an endogenous agonist for a C. elegans TRPV OSM-9/OCR-4 channel. Nature Communications, 7, 1–11. https://doi.org/10.1038/ncomms13135 Vamos, E., Pardutz, A., Klivenyi, P., Toldi, J., & Vecsei, L. (2009). The role of kynurenines in disorders of the central nervous system: Possibilities for neuroprotection. Journal of the Neurological Sciences, 283(1–2), 21–27. https://doi.org/10.1016/j.jns.2009.02.326 Van Beijnum, J. R., Moerkerk, P. T. M., Gerbers, A. J., De Brune, A. P., Arends, J. W., Hoogenboom, H. R., & Hufton, S. E. (2002). Target validation for genomics using peptide- specific phage antibodies: A study of five gene products overexpressed in colorectal cancer. International Journal of Cancer, 101(2), 118–127. https://doi.org/10.1002/ijc.10584 Vrablik, T. L., Huang, L., Lange, S. E., & Hanna-Rose, W. (2009). Nicotinamidase modulation of NAD+ biosynthesis and nicotinamide levels separately affect reproductive development and cell survival in C. elegans. Development, 136(21), 3637–3646. https://doi.org/136/21/3637 [pii]10.1242/dev.028431 Vrablik, T. L. T. L., Huang, L., Lange, S. E. S. E., & Hanna-Rose, W. (2009). Nicotinamidase 108

modulation of NAD+ biosynthesis and nicotinamide levels separately affect reproductive development and cell survival in C. elegans. Development (Cambridge, England), 136(21), 3637–46. https://doi.org/10.1242/dev.028431 Vrablik, T. L., Wang, W., Upadhyay, A., & Hanna-Rose, W. (2011). Muscle type-specific responses to NAD+ salvage biosynthesis promote muscle function in Caenorhabditis elegans. Developmental Biology, 349(2), 387–94. https://doi.org/10.1016/j.ydbio.2010.11.014 Wang, B., Hasan, M. K., Alvarado, E., Yuan, H., Wu, H., & Chen, W. Y. (2011). NAMPT overexpression in prostate cancer and its contribution to tumor cell survival and stress response. Oncogene, 30(8), 907–921. https://doi.org/10.1038/onc.2010.468 Wang, L., Ding, D., Salvi, R., & Roth, J. A. (2014). Nicotinamide adenine dinucleotide prevents neuroaxonal degeneration induced by manganese in cochlear organotypic cultures. NeuroToxicology, 40, 65–74. https://doi.org/10.1016/j.neuro.2013.11.007 Wang, W., Mcreynolds, M. R., Goncalves, J. F., Shu, M., Dhondt, I., Braeckman, B. P., … Hanna-rose, X. W. (2015). Comparative Metabolomic Profiling Reveals That Dysregulated Glycolysis Stemming from Lack of Salvage NAD Biosynthesis Impairs Reproductive Development in Caenorhabditis elegans * □, 290(43), 26163–26179. https://doi.org/10.1074/jbc.M115.662916 Watson, M., Roulston, A., Bélec, L., Billot, X., Marcellus, R., Bédard, D., … Beauparlant, P. (2009). The small molecule GMX1778 is a potent inhibitor of NAD+ biosynthesis: strategy for enhanced therapy in nicotinic acid phosphoribosyltransferase 1-deficient tumors. Molecular and Cellular Biology, 29(21), 5872–88. https://doi.org/10.1128/MCB.00112-09 Williams, B., Jones, P., & Agarose, R. (1985). Calmodulin-Dependent NAD Kinase of Human Neutrophils has been partially purified by sequential application activity observed at free calcium concentrations of activation of NAD kinase increases the maximum velocity of the reaction from 2- to 5-fold while , 237(1), 80–87. Xu, P., & Sauve, A. A. (2010). Vitamin B3, the nicotinamide adenine dinucleotides and aging. Mechanisms of Ageing and Development, 131(4), 287–298. https://doi.org/10.1016/j.mad.2010.03.006 Yang, H., Lavu, S., & Sinclair, D. A. (2006). Nampt/PBEF/Visfatin: A regulator of mammalian health and longevity? Experimental Gerontology, 41(8), 718–726. https://doi.org/10.1016/j.exger.2006.06.003 Yang, T., Chan, N. Y. K., & Sauve, A. A. (2007). Syntheses of nicotinamide riboside and derivatives: Effective agents for increasing nicotinamide adenine dinucleotide concentrations in mammalian cells. Journal of Medicinal Chemistry, 50(26), 6458–6461. https://doi.org/10.1021/jm701001c Yang, Y., & Sauve, A. A. (2016). NAD+ metabolism: Bioenergetics, signaling and manipulation for therapy. Biochimica et Biophysica Acta - Proteins and Proteomics, 1864(12), 1787– 1800. https://doi.org/10.1016/j.bbapap.2016.06.014 Yoshino, J., Mills, K. F., Yoon, M. J., & Imai, S. I. (2011). Nicotinamide mononucleotide, a key NAD + intermediate, treats the pathophysiology of diet- and age-induced diabetes in mice. Cell Metabolism, 14(4), 528–536. https://doi.org/10.1016/j.cmet.2011.08.014 Zhang, H., Ryu, D., Wu, Y., Gariani, K., Wang, X., Luan, P., … Auwerx, J. (2016). Supplementary Materials for enhances life span in mice. Science, 6(6292), 1436–1443. https://doi.org/10.1126/science.aaf2693 Zhou, M., Ottenberg, G., Sferrazza, G. F., Hubbs, C., Fallahi, M., Rumbaugh, G., … Lasmezas, 109

C. (2015). Neuronal death induced by misfolded prion protein is due to NAD+ depletion and can be relieved in vitro and in vivo by NAD+ replenishment. Brain : A Journal of Neurology, 138(Pt 4), 992–1008. https://doi.org/10.1093/brain/awv002

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Appendix: Amino acids are not used as an energy source in pnc-1 mutants

We previously reported that NAD+ salvage synthesis through the nicotinamidase PNC-1 is required for normal progression of gonad development in C. elegans. Global metabolic profiling suggested that glycolysis was perturbed in our pnc-1 mutants, which have lower global levels of NAD+. I used metabolic tracing analysis in wild type and pnc-1 mutants to confirm that glycolysis is compromised when NAD+ salvage synthesis is blocked. Although glycolysis is impaired, we had evidence that mitochondrial functions were not changed in pnc-1.

Metabolomics analysis showed that unlike other glycolytic intermediates pyruvate levels are not reduced in pnc-1 mutants. Therefore, we initially hypothesized that excessive use of amino acids as an energy source compensates for insufficient glycolytic flux in pnc-1 mutants. Using a targeted metabolomics approach, I showed that alanine steady state levels are increased 2-fold in our pnc-1 mutants (Figure A-1). In addition to this, α-ketoglutarate and glutamate steady state levels are also significantly elevated in pnc-1 mutants compared to WT (Figure A-2). This data suggested that a steady reservoir of amino acids is in place for pyruvate production through protein degradation due to glycolytic blockage. I also observed an up-regulation of mRNA expression of an important alanine aminotransferase, agxt-1, in our pnc-1 mutants; supporting the notion that amino acid catabolism is indeed occurring (Figure A-3). Our results suggest that compromised glycolysis activates amino acid catabolism to maintain functions of the mitochondria. However, this metabolic shift is not compatible with normal progression of reproductive development in C. elegans. To further investigate this hypothesis, I used RNAi to knockdown agxt-1 in wild-type animals and pnc-1 mutants. I predicted that pyruvate steady state levels would decrease in pnc-1 mutants treated with agxt-1 RNAi. However, compared to controls pyruvate levels did not change when agxt-1 was knockdown in pnc-1 mutants (Figure 111

A-4). Also, I used metabolic tracing analysis to investigate this hypothesis. I predicted that we would observe an increase of isotope label from alanine being incorporated into pyruvate in pnc-

1 mutants compared to wild-type animals. However, this was not the case. After five hours of treating C. elegans cultures with isotope alanine, we detected a decrease in label entering pyruvate in our pnc-1 mutants (Figure A-5). Therefore, if amino acids are compensating for an insufficient lack of glycolytic flux remains inconclusive. Based on our results, there is a compensatory mechanism occurring the supply carbon to the mitochondria for oxidative

phosphorylation, but we are unable to conclude that it is

amino acid catabolism.

Figure A-1: Alanine steady state levels are increased in pnc-1 mutants. Shown are LC-MS measurements of global Alanine levels in N2 and pnc-1(pK9605) mutants cultured on UV-killed OP50. Boxes show the upper and lower quartile values, + indicates the mean value and lines indicate the median. Error bars indicate the maximum and minimum of the population distribution. *, 0.01

Figure A-2: α-ketoglutarate and glutamate steady state levels are increased in pnc-1 mutants. Shown are LC-MS measurements of global α-ketoglutarate and glutamate levels in N2 and pnc- 1(pK9605) mutants cultured on UV-killed OP50. Boxes show the upper and lower quartile values, + indicates the mean value and lines indicate the median. Error bars indicate the maximum and minimum of the population distribution. *, 0.01

Figure A-3: Alanine aminotransferase mRNA levels are up-regulated in pnc-1 mutants. qRT-PCR analysis of agx-1 mRNA levels in pnc-1 mutants compared to N2 on UV-killed OP50. *, 0.01

Figure A-4: Pyruvate steady state levels in worms treated with agxt-1 RNAi. Shown are LC-MS measurements of normalized pyruvate peak area in N2 and pnc-1(pK9605) mutants cultured on UV-killed OP50. 113

Figure A-5: Metabolic carbon tracing with stable isotope alanine in N2 and pnc-1 mutants. 13 Shown are LC-MS measurements of (A) % incorporation of 3C -Alanine in N2 and pnc-1 mutants. Dot- plot represents each biological replicate in wild-type and pnc-1(pK9605) mutants after 3 and 5 hours of 13 13 exposure to 3C -Alanine (B) % Incorporation of 3C -Pyruvate in wild-type and pnc-1 mutants after 3 and 13 5 hours of exposure to 3C -Alanine.

VITA

Melanie R. McReynolds Education • 2011-Present, Ph.D. in Biochemistry, Microbiology and Molecular Biology, The Pennsylvania State University • 2009-2011, M.S. (Highest Honors) in Biological Sciences, Alcorn State to Penn State Bridges to the Doctorate Program, Alcorn State University • 2005-2009, B.S.(Magna Cum Laude) in Chemistry and Physics, Alcorn State University

Publications • Crook, M., McReynolds, M., Wang, W., Hanna-Rose, W. (2014). An NAD+ Biosynthetic Pathway Enzyme Functions Cell Non-Autonomously in C. elegans Development. Developmental Dynamics. 243:965-967. • Wang W., McReynolds M.R., Gonvalves, J.F., Shu, M., Dhondt, I., Braeckman, B.P., Lange, S.E., Kho, K. Detwiler, A.C., Pacella, M.J. and W. Hanna-Rose. (2015). Comparative metabolomic profiling reveals that dysregulated glycolysis stemming from lack of salvage NAD+ biosynthesis impairs reproductive development in C. elegans J. Bio. Chem. 2015, 290:26163-26179. • Ozcelik, A., Nama, N., Huang, PH., Kaynak, M., McReynolds, MR., Hanna-Rose, W., Huang, TJ. (2016). Acoustofluidic rotational manipulation of cells and organisms using oscillating solid structures. SMALL. DOI: 10.1002/smll.201601760 • McReynolds, M.R., Wang, W., Holleran, L.M., Hanna-Rose, W. (2017). Uridine monophosphate synthetase 1 enables eukaryotic de novo NAD+ biosynthesis from quinolinic acid. In review

Awards and Honors • 2017: University Student Way Paver Award—Council College of Multicultural Leaders (CCML), Penn State University • 2016: FASEB MARC Travel Award- Postdoctoral Preparation Institute Workshop • 2016: ASBMB 2016 Best Thematic Poster Award- Metabolism, Disease and Drug Design • 2016: St. Jude National Graduate Student Symposium (NGSS)—Invited/Selected Participant • 2016: ASBMB MAC Travel Award- Experimental Biology meeting • 2015: FASEB MARC Travel Award- FASEB Grant Writing Seminar & Responsible Conduct of Research Workshop • 2015: FASEB MARC Program Poster/Oral Presentation Travel Award- GSA: 20th International C. elegans meeting • 2014: Alfred P. Sloan MPHD Scholar, Penn State University • 2012: Bunton-Waller Fellowship, Penn State University