A thesis submitted in fulfilment of the requirements for the
degree of Doctor of Philosophy
Metabolic tracing of NAD+ precursors using
strategically labelled isotopes of NMN
by
Lynn-Jee Kim
Supervisors
Dr. Lindsay E. Wu (Primary)
Prof. David A. Sinclair (Joint-Primary)
Dr. Lake-Ee Quek (Co-supervisor)
School of Medical Sciences
Faculty of Medicine
Thesis/Dissertation Sheet
Surname/Family Name: Kim Given Name/s: Lynn-Jee Abbreviation for degree as give in PhD the University calendar: Faculty: Faculty of Medicine School: School of Medical Sciences + Thesis Title: Metabolic tracing of NAD precursors using strategically labelled isotopes of NMN
Abstract 350 words maximum: (PLEASE TYPE) Nicotinamide adenine dinucleotide (NAD+) is an important cofactor and substrate for hundreds of cellular processes involved in redox homeostasis, DNA damage repair and the stress response. NAD+ declines with biological ageing and in age-related diseases such as diabetes and strategies to restore intracellular NAD+ levels are emerging as a promising strategy to protect against metabolic dysfunction, treat age-related conditions and promote healthspan and longevity. One of the most effective ways to increase NAD+ is through pharmacological supplementation with NAD+ precursors such as nicotinamide mononucleotide (NMN) which can be orally delivered. Long term administration of NMN in mice mitigates age-related physiological decline and alleviates the pathophysiologies associated with a high fat diet- and age-induced diabetes. Despite such efforts, there are certain aspects of NMN metabolism that are poorly understood. In this thesis, the mechanisms involved in the utilisation and transport of orally administered NMN were investigated using strategically labelled isotopes of NMN and mass spectrometry. A mass spectrometry method was developed to trace the incorporation of labelled NMN moieties in NAD+ metabolites following supplementation with labelled NMN compounds. This was validated in biologically relevant models such as mammalian cell lines (Chapter 3) and bacteria (Chapter 4), the latter serving as a proof-of-concept model to investigate NMN metabolism through bacterial routes before investigating its metabolic fate in vivo (Chapter 5). Following oral administration with labelled NMN compounds in mice, labelled NAD+ metabolites were detected in abundance in the peripheral tissues of mice treated with antibiotics but were largely absent in control mice. This suggests the majority of orally administered NMN is consumed by gut bacteria, limiting its availability to host peripheral tissues and insinuates host-microbe competition for NAD+ precursors. Interestingly, an abundance of nicotinamide riboside (NR) was observed both in vitro and in vivo following supplementation with NMN supporting the indirect NMN transport mechanism whereby it is dephosphorylated to NR prior to entering the cell. Overall, these findings have therapeutic implications in the dosing and route of administration of NMN as an NAD+-boosting strategy to treat conditions related to metabolic dysfunction and age-related diseases and further to promote healthy ageing and longevity.
Declaration relating to disposition of project thesis/dissertation
I hereby grant to the University of New South Wales or its agents the right to archive and to make available my thesis or dissertation in whole or in part in the University libraries in all forms of media, now or here after known, subject to the provisions of the Copyright Act 1968. I retain all property rights, such as patent rights. I also retain the right to use in future works (such as articles or books) all or part of this thesis or dissertation.
I also authorise University Microfilms to use the 350 word abstract of my thesis in Dissertation Abstracts International (this is applicable to doctoral theses only).
…………………………………… …………………………… ………...… Signature Witness Signature Date The University recognises that there may be exceptional circumstances requiring restrictions on copying or conditions on use. Requests for restriction for a period of up to 2 years must be made in writing. Requests for a longer period of restriction may be considered in exceptional circumstances and require the approval of the Dean of Graduate Research.
FOR OFFICE USE ONLY Date of completion of requirements for Award:
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INCLUSION OF PUBLICATIONS STATEMENT
UNSW is supportive of candidates publishing their research results during their candidature as detailed in the UNSW Thesis Examination Procedure.
Publications can be used in their thesis in lieu of a Chapter if: • The candidate contributed greater than 50% of the content in the publication and is the “primary author”, ie. the candidate was responsible primarily for the planning, execution and preparation of the work for publication • The candidate has approval to include the publication in their thesis in lieu of a Chapter from their supervisor and Postgraduate Coordinator. • The publication is not subject to any obligations or contractual agreements with a third party that would constrain its inclusion in the thesis
Please indicate whether this thesis contains published material or not: This thesis contains no publications, either published or submitted for ☐ publication (if this box is checked, you may delete all the material on page 2)
Some of the work described in this thesis has been published and it has ☒ been documented in the relevant Chapters with acknowledgement (if this box is checked, you may delete all the material on page 2)
This thesis has publications (either published or submitted for publication) ☐ incorporated into it in lieu of a chapter and the details are presented below
CANDIDATE’S DECLARATION I declare that: • I have complied with the UNSW Thesis Examination Procedure • where I have used a publication in lieu of a Chapter, the listed publication(s) below meet(s) the requirements to be included in the thesis. Candidate’s Name Signature Date (dd/mm/yy) Lynn-Jee Kim
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Acknowledgements
First and foremost, I would like to sincerely thank my supervisors, Lindsay, David and Lake- Ee for their support and for providing me with all the necessary means and opportunities during my PhD. I thank each of you for all the intellectual insight, genuine advice and guidance I have received over the years. Thank you for all the invaluable memories, both within and outside the lab and for always making the extra effort to make the research experience in our lab unique and inspiring.
A special thank you to all the members, past and present, of the Laboratory for Ageing Research. Thank you for being the most amazing team to work with, as well as learn alongside and from. Thank you especially to Abhi, Catherine, Jon, Ashley, Jin, and Tim for their assistance in experiments and for helping me solve experimental and technical dilemmas. This gratitude is also extended to faculty members and fellow peers of 3East, Allison and BRC technical staff for all their guidance and support.
Support as part of the Australian Post-Graduate award is kindly acknowledged and I sincerely thank the post-graduate research support scheme for funding domestic and international conference travels.
Finally, to all my friends and family, I thank you all for the undying love and encouragement I have received throughout my PhD. Here’s to growing older and healthier together.
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Publications and Presentations Research Publications
NAD+ Repletion Rescues Female Fertility During Reproductive Ageing. Cell Reports, Feb 2020 Michael J. Bertoldo, Dave R. Listijono, Wing-Hong Jonathan Ho, Angelique H. Riepsamen, Xing L. Jin, Kaisa Selesniemi, Dale M. Goss, Saabah Mahbub, Jared M. Campbell, Abbas Habibalahi, Wei-Guo Nicholas Loh, Neil A. Youngson, Jayanthi Maniam, Ashley S.A. Wong, Dulama Richani, Catherine Li, Yiqing Zhao, Maria Marinova, Lynn-Jee Kim, Laurin Lau, Rachael M Wu, A. Stefanie Mikolaizak, Toshiyuki Araki, David G. Le Couteur, Nigel Turner, Margaret J. Morris, Kirsty A. Walters, Ewa Goldys, Christopher O’Neill, Robert B. Gilchrist, David A. Sinclair, Hayden A. Homer, Lindsay E. Wu
+ Impairment of An Endothelial NAD -H2S Signalling Network Is A Reversible Cause Of Vascular Aging. Cell, Mar 2018. Abhirup Das, George X. Huang, Michael S. Bonkowski, Alban Longchamp, Catherine Li, Michael B. Schultz, Lynn-Jee Kim, Brenna Osborne, Sanket Joshi, Yuancheng Lu, Jose Humberto Treviño-Villarreal, Myung-Jin Kang, Tzong-tyng Hung, Brendan Lee, Eric O. Williams, Masaki Igarashi, James R. Mitchell, Lindsay E. Wu, Nigel Turner, Zolt Arany, Leonard Guarente, David A. Sinclair
Nicotinamide mononucleotide (NMN) deamidation and indirect regulation of the NAD metabolome Submitted to Nature Communications on 02/10/2020 Lynn-Jee Kim, Timothy J. Chalmers, Greg C. Smith, Abhirup Das, Eric Wing Keung Poon, Jun Wang, Simon P. Tucker, David A. Sinclair, Lake-Ee Quek, Lindsay E. Wu
Fertility preservation during chemotherapy treatment without compromising oncological efficacy by NAD+ repletion. In preparation for resubmission Wing-Hong Jonathan Ho, Dave R. Listijono, Michael J. Bertoldo, Angelique H. Riepsamen, Kaisa Selesniemi, Yiqing Zhao, Wei-Guo Nicholas Loh, Neil A. Youngson, Safaa Cabot,
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Ashley S.A. Wong, Dulama Richani, Catherine Li, Lynn-Jee Kim, Laurin Lau, Rachael M Wu, Pawel Kordowitzki, Toshiyuki Araki, A. Stefanie Mikolaizak, Sonia Bustamante, Abhirup Das, Jayanthi Maniam, David G. Le Couteur, Nigel Turner, Lake-Ee Quek, Margaret J. Morris, Kirsty A. Walters, Robert B. Gilchrist, David A. Sinclair, Hayden A. Homer, Lindsay E. Wu
Oral Presentations
Metabolic Tracing of NAD+ Precursors Using Strategically Labelled Isotopes of NMN: Contribution of The Gut Microbiome to NAD+ Metabolism. Lynn-Jee Kim, Jonathan Ho, Jo Ann Yap, Lake-Ee Quek, David A. Sinclair, Lindsay E. Wu Neuroscience and Non-Communicable Diseases Department Seminar 2019, UNSW, Australia
Investigating Novel NAD+ Precursors and Their Therapeutic Potential Against Age- Associated Physiological Decline. (Poster selected for oral presentation) Lynn-Jee Kim, Lake-Ee Quek, David A. Sinclair, Lindsay E. Wu IUBMB Seoul 2018 Meeting, COEX Convention Centre 2018, Seoul, South Korea.
Investigating Novel NAD+ Precursors and Their Therapeutic Potential Against Age- Associated Physiological Decline. Lynn-Jee Kim, Lake-Ee Quek, David A. Sinclair, Lindsay E. Wu Neuroscience and Non-Communicable Diseases Department Seminar 2017, UNSW, Australia
Poster Presentations
Gut Microbiome Competes with Host For Orally Administered Nicotinamide Mononucleotide (NMN) By Reducing Its Availability And Altering Uptake Pathways. Lynn-Jee Kim, Jonathan Ho, Jo Ann Yap, Lake-Ee Quek, David A. Sinclair, Lindsay E. Wu Australian Biology of Ageing Conference 2019, University of Sydney, NSW, Australia. Awarded Best Poster Presentation.
Investigating Novel NAD+ Precursors and Their Therapeutic Potential Against Age- Associated Physiological Decline. Lynn-Jee Kim, Lake-Ee Quek, David A. Sinclair, Lindsay E. Wu
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IUBMB Seoul 2018 Conference, COEX Convention Centre, Seoul, South Korea. Awarded Top 10 Best Poster Presentations
Investigating Novel NAD+ Precursors and Their Therapeutic Potential Against Age- Associated Physiological Decline. Lynn-Jee Kim, Lake-Ee Quek, David A. Sinclair, Lindsay E. Wu Australian Biology of Ageing Conference 2018, University of Queensland, Queensland, Australia
Metabolic Tracing of NAD+ Precursors Using Strategically Labelled Isotopes. Lynn-Jee Kim, Lake-Ee Quek, David A. Sinclair, Lindsay E. Wu Cell Symposia Meeting: Aging and Metabolism 2018, Sitges, Spain.
Metabolic Tracing of NAD+ Precursors Using Strategically Labelled Isotopes. Lynn-Jee Kim, Lake-Ee Quek, David A. Sinclair, Lindsay E. Wu Cold Spring Harbor Laboratory 2018 Meeting: Mechanisms of Aging, Cold Spring Harbor, New York, USA.
The Role of NAD+ Biology in Chemotherapy-Induced Cardiotoxicity. Lynn-Jee Kim, Abhirup Das, David A. Sinclair, Lindsay E. Wu Australian Biology of Ageing Conference 2016, Coogee, NSW, Australia
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Table of Contents
Acknowledgements ...... vi Publications and Presentations ...... vii Abstract ...... xiii List of Figures ...... xiv List of Tables ...... xvi List of Abbreviations ...... xvii ...... 1 1.1 Adding Health to Years ...... 1 1.2 The Biological Significance of NAD+ ...... 5 1.2.1 NAD+ as a redox cofactor ...... 8 1.2.2 NAD+ as a substrate ...... 15 1.3 NAD+ deficiency is a critical pathological factor in ageing and disease ...... 39 1.3.1 NAD+ deficiency in ageing and disease ...... 39 1.3.2 Increased NAD+ consumption as a cause of NAD+ deficiency ...... 40 1.3.3 Decreased NAD+ biosynthesis as a cause of NAD+ deficiency ...... 42 1.4 Consequences of NAD+ deficiency in cell function and survival ...... 44 1.5 Therapeutic potential of NAD+ boosting strategies ...... 46 1.6 NAD+ biosynthesis pathways ...... 49 1.6.1 The de novo synthesis pathway ...... 49 1.6.2 The Preiss-Handler pathway ...... 50 1.6.3 The Salvage/Recycling pathway ...... 51 1.6.4 Additional pathways in yeast and bacteria, that do not exist in mammals ...... 53 1.7 Current gaps in the field of NAD+ biology ...... 55 1.7.1 Unexplored mechanisms of NAD+ precursor deamidation ...... 55 1.7.2 NAD+ precursor uptake mechanisms still unclear ...... 56 1.7.3 Quantifying the NAD+ metabolome remains a challenge ...... 57 1.8 Thesis goals ...... 59 ...... 61 2.1 Isotopic labelled compounds...... 61 2.1.1 Synthesis of labelled NMN compounds ...... 61 2.1.2 Labelled glutamine...... 63 2.2 Sample processing for mass spectrometry ...... 64 2.2.1 Preparation of extraction buffer ...... 64
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2.2.2 Cell harvest and extraction...... 65 2.2.3 Further sample processing for mass spectrometry ...... 65 2.2.4 NAD+ metabolite standard curve preparation ...... 65 2.3 LC-MS/MS ...... 66 2.3.1 Liquid chromatography ...... 66 2.3.2 Mass spectrometry ...... 67 2.4 Data analysis ...... 67 2.5 Statistics ...... 68 ...... 69 3.1 Introduction ...... 69 3.2 Methods...... 77 3.2.1 Optimisation of mass spectrometry parameters ...... 77 3.2.2 Liquid chromatography (LC) separation ...... 78 3.2.3 Isolation of primary hepatocytes and treatment ...... 79 3.2.4 C2C12 and HEK293 Cell Culture...... 81 3.3 Results ...... 82 3.3.1 Compound optimisation of mass spectrometry parameters for internal standards. 84 3.3.2 Optimisation of mass spectrometry parameters for NAD+ metabolites ...... 85 3.3.3 Detecting labelled NMN isotopes using LC-MS/MS and MRM...... 89 3.3.4 Tracing the metabolic fate of NMN isotopes using LC-MS/MS to detect labelled NAD+ metabolites...... 90 3.3.5 Intracellular NR is detected following extracellular supplementation of NMN in primary hepatocytes...... 95 3.3.6 NMN does not lead to increases in deamidated metabolites, NaMN and NaAD, following supplementation in primary hepatocytes...... 98 3.3.7 Time-course tracing of NMN metabolism in C2C12 and HEK293 cells – amidated metabolites ...... 106 3.3.8 Time-course tracing of NMN metabolism in C2C12 and HEK293 cells – deamidated metabolites ...... 110 3.4 Discussion ...... 115 ...... 121 4.1 Introduction ...... 121 4.1.1 Bacterial de novo NAD biosynthesis pathway...... 123 4.1.2 NAD+ is salvaged by deamidation in bacteria ...... 125 4.1.3 Potential NMN uptake routes in bacteria ...... 127 4.1.4 Key differences compared to eukaryotic pathway ...... 129
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4.2 Methods...... 131 4.2.1 Bacterial culture and NMN treatment ...... 131 4.2.2 Extracting NAD+ metabolites from E. coli cell lysates and culture supernatants. 134 4.3 Results ...... 139 4.3.1 Supplementation of NMN in E. coli – changes to NAD+ metabolites...... 139 4.4 Discussion ...... 147 ...... 152 5.1 Introduction ...... 152 5.1.1 Rationale for NMN isotope design ...... 157 5.2 Methods...... 164 5.2.1 Animal housing ...... 164 5.2.2 Antibiotic and NMN treatment regimen ...... 164 5.2.3 Plasma collection and metabolite extraction...... 165 5.2.4 Tissue collection, metabolite extraction and data analysis ...... 167 5.2.5 DNA extraction from faecal pellets ...... 168 5.3 Results ...... 170 5.3.1 Antibiotic ablation of the gut microbiome ...... 170 5.3.2 Antibiotic treatment increases NAD+ metabolite and NMN availability in the intestinal tissue of mice...... 176 5.3.3 Evidence for NMN uptake facilitated by dephosphorylation into NR ...... 186 5.3.4 NMN deamidation is independent of the microbiome...... 190 5.4 Discussion ...... 193 ...... 201 6.1 Quantifying the NAD metabolome using LC-MS/MS—So close, yet [not] so far . 203 6.2 Mechanisms of NMN uptake in gut epithelial tissue — ¿Por qué no los dos? ...... 206 6.3 Host-microbe interactions in NAD+ precursor metabolism—Friend or foe? ...... 208 6.4 Preliminary evidence for host NMN deamidation—it’s not you, it’s me ...... 210 6.5 Beyond NAD metabolism—blue-sky thinking ...... 211 Appendix ...... 213 References ...... 281
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Abstract
Nicotinamide adenine dinucleotide (NAD+) is an important cofactor and substrate for hundreds of cellular processes involved in redox homeostasis, DNA damage repair and the stress response. NAD+ declines with biological ageing and in age-related diseases. Strategies to restore intracellular NAD+ levels are emerging as promising therapeutics to protect against metabolic dysfunction, treat age-related conditions and promote healthspan and longevity. One of the most effective ways to increase NAD+ is through pharmacological supplementation with NAD+ precursors such as nicotinamide mononucleotide (NMN), which can be orally delivered. Long term administration of NMN in mice mitigates age-related physiological decline and alleviates pathophysiologies associated with a high fat diet- and age-induced diabetes. Despite such efforts, there are certain aspects of NMN metabolism that are poorly understood. In this thesis, the mechanisms involved in the utilisation and transport of orally administered NMN were investigated using strategically labelled isotopes of NMN and mass spectrometry. A mass spectrometry method was developed to trace the incorporation of labelled NMN moieties into NAD+ metabolites following supplementation with labelled NMN compounds. This was validated in biologically relevant models such as mammalian cell lines (Chapter 3) and bacteria (Chapter 4), the latter serving as a proof-of-concept model to investigate NMN metabolism through bacterial routes before investigating its metabolic fate in vivo (Chapter 5). Following oral administration with labelled NMN compounds in mice, labelled NAD+ metabolites were detected in abundance in the peripheral tissues of mice treated with antibiotics but were largely absent in control mice. This suggests the majority of orally administered NMN is consumed by gut bacteria, limiting its availability to host peripheral tissues and insinuates host-microbe competition for NAD+ precursors. Interestingly, an abundance of nicotinamide riboside (NR) was observed both in vitro and in vivo following supplementation with NMN, supporting the indirect NMN transport mechanism whereby it is dephosphorylated to NR prior to entering the cell. Overall, these findings have therapeutic implications in the dosing and route of administration of NMN as an NAD+-boosting strategy to treat conditions related to metabolic dysfunction and age-related diseases and further to promote healthy ageing and longevity.
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List of Figures Figure 1.1 Chemical structure of NAD+ ...... 5 Figure 1.2 The biological roles of NAD+ ...... 7 Figure 1.3 Functions of NAD(P)(H) as a redox cofactor...... 14 Figure 1.4 NAD+ biosynthesis pathways in mammals...... 54 Figure 2.1 Chemical structure and isotope labelling positions of NMN ...... 62 Figure 2.2 Chromatogram peaks of NMN isotopes using LC-MS/MS injected at 100 µM. ... 63 Figure 2.3 Chemical structure and isotope labelling of glutamine...... 63 Figure 2.4 Chromatogram peaks of unlabelled glutamine and isotope labelled glutamine using LC-MS/MS injected at 100 µM...... 64 Figure 3.1 Labelling strategy of labelled NMN isotopes...... 74 Figure 3.2 Detecting labelled NMN and NAD+ isotopologues using MRM (multiple reaction monitoring)...... 83 Figure 3.3 Representative chromatograms of unlabelled (M+0) and labelled NMN isotopes from injection of a 100 µM standard solution...... 89 Figure 3.4 Labelled NMN compounds are detected intact and contribute to an increase in NAD+ in primary hepatocytes...... 93 Figure 3.5 Labelled NAD+ is degraded and contributes to increases in metabolites in the NAD+ recycling pathway...... 94 Figure 3.6 An abundance of intact labelled NR is detected following supplementation with labelled NMN...... 97 Figure 3.7 Chromatogram peaks of NMN/NaMN isotopologue pairs sharing the same MRM transition...... 102 Figure 3.8 Chromatogram peaks from NAD+/NaAD isotopologues sharing the same MRM transition...... 105 Figure 3.9 Labelled NMN detected intact and contributes to increases in NAD+ and NR over time in cell...... 108 Figure 3.10 Degradation of NAD+ and nicotinamide salvage ...... 109 Figure 3.11 Detection of NMN/NaMN isotopologue pairs in C2C12 and HEK293 cells following supplementation with NMN...... 113 Figure 3.12 Detection of NAD+/NaAD isotopologue pairs in C2C12 and HEK293 cells following supplementation with NMN...... 114 Figure 4.1 NAD+ biosynthesis pathways in bacteria...... 126 Figure 4.2 E. coli growth curve and sample OD600 values...... 132 Figure 4.3 E. coli growth curve and sample OD600 values...... 133 Figure 4.4 Detection of amidated NAD+ metabolites in the supernatant and cell lysate from E. coli following supplementation with NMN...... 143
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Figure 4.5 Detection of amidated NAD+ metabolites in the supernatant and cell lysate from E. coli following supplementation with NMN1...... 144 Figure 4.6 Detection of deamidated NAD+ metabolites in the supernatant and cell lysate from E. coli following supplementation with NMN...... 145 Figure 4.7 Detection of deamidated NAD+ metabolites in the supernatant and cell lysate from E. coli following supplementation with NMN1...... 146 Figure 5.1 Labelled isotope compounds ...... 158 Figure 5.2 Schematic of the NMN deamidation and re-amidation hypothesis using NMN1 and 15N-glutamine...... 161 Figure 5.3 Schematic of NMN deamidation hypothesis using NMN2...... 162 Figure 5.4 Schematic for the potential biosynthesis pathways and NAD+ metabolites involved in the utilisation of NMN2...... 163 Figure 5.5 The effect of antibiotic treatment in mice...... 173 Figure 5.6 Reduced diversity of the murine faecal microbiome upon antibiotic treatment in NMN2 cohort...... 174 Figure 5.7 Reduced diversity of the murine faecal microbiome upon antibiotic treatment in NMN1 cohort...... 175 Figure 5.8 An increase in amidated NAD+ metabolites detected with antibiotic treatment in the intestinal tissue after NMN2 administration...... 180 Figure 5.9 Detection of amidated NAD+ metabolites with antibiotic treatment in the liver after NMN2 administration...... 181 Figure 5.10 Detection of nicotinamide (NAM) with antibiotic treatment in the plasma after NMN2 administration...... 182 Figure 5.11 An increase in amidated NAD+ metabolites detected with antibiotic treatment in the intestinal tissue after NMN1 administration...... 183 Figure 5.12 Detection of amidated NAD+ metabolites with antibiotic treatment in the liver after NMN1 administration...... 184 Figure 5.13 Detection of nicotinamide (NAM) with antibiotic treatment in the plasma after NMN1 administration...... 185 Figure 5.14 Schematic for the direct or indirect transport of NMN in intestinal cells...... 188 Figure 5.15 The ratio of the labelled-to-unlabelled isotopologues of NAD+ metabolites ..... 189 Figure 5.16 Deamidation pathway in the intestinal tissue of antibiotic treated mice following administration with NMN2 or NMN1 and 15N-amide-glutamine (15N-gln)...... 192
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List of Tables Table 3.1 Liquid chromatography (LC) separation gradient ...... 78 Table 3.2 MRM transitions for internal standards and mass spectrometry parameters...... 84 Table 3.3 MRM transitions and MS parameters for unlabelled NAD+ metabolites ...... 85 Table 3.4 MRM transitions and mass spectrometry parameters for isotopologues of NAM, NMN, NR and NAD+ ...... 86 Table 3.5 MRM transitions and mass spectrometry parameters for isotopologues of NA, NaR, NaMN and NaAD ...... 87 Table 3.6 LC-MS precision and accuracy parameters for each metabolite ...... 88 Table 4.1 MRM transitions and mass spectrometry (MS) parameters to detect NAD+ metabolites in E. coli (supernatant and lysates) following supplementation with unlabelled NMN using LC-MS/MS...... 136 Table 4.2 MRM transitions and MS parameters to detect NAD+ isotopologues of amidated metabolites in E. coli (supernatant and lysates) following supplementation with NMN1 using LC-MS/MS...... 137 Table 4.3 MRM transitions and MS parameters to detect NAD+ isotopologues of deamidated metabolites in E. coli (supernatant and lysates) following supplementation with NMN1 using LC-MS/MS...... 138
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Chapter 1 Introduction
List of Abbreviations
3-HAO 3-hydroxyanthranolic acid oxygenase 6-PG 6-phosphogluconate Abx Antibiotics ACN Acetonitrile AD Alzheimer's Disease ADP Adenosine diphosphate ADPR Adenosine diphosphate ribose AK Adenosine kinase ALC1 Amplified in liver cancer protein 1 aNHEJ Alternative pathway for non-homologous end joining AOPCP Adenosine-5′-(α,β-methylene) diphosphate ART Adenosine diphosphate-ribosyltransferase ATP Adenosine triphosphate BER Base excision repair BRCA1 Breast cancer type 1 susceptibility protein BRCT BRCA1 C-terminus BRMS1 Breast cancer metastasis suppressor 1 BSA Bovine serum albumin C2C12 Mouse myoblast cell line cADPR Cyclic adenosine diphosphate ribose CD157 Cluster of differentiation 157 CD38 Cluster of differentiation 38 CD73 Cluster of differentiation 73 CE Collision energy CID Collision induced dissociation CK2 Casein kinase 2 cNHEJ Classical pathway for non-homologous end joining CobT Cobaltochelatase CPS1 Carbamoyl phosphate synthetase 1 CS Cockayne Syndrome CSA Camphorsulfonic acid (internal standard) CtBP C-terminal binding protein CtIP C-terminal interacting protein CXP Cell exit potential cyclin-B/cdk1 Cyclin B/cyclin-dependent kinase 1 CYP450 Cytochrome P450 DBC1 Deleted in breast cancer 1 DDB2 DNA damage-binding protein 2 DDR DNA damage response DHAP Dihydroxyacetone phosphate Dip Dipyridamole
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Chapter 1 Introduction
DMEM Dulbecco's minimum essential medium DNA-PKcs DNA-dependent protein kinase catalytic subunit DP Declustering potential DSB Double-stranded breaks DYRK Dual specificity tyrosine phosphorylation-regulated kinases EGTA Ethylene glycol-bis(β-aminoethyl ether)-N,N,N′,N′- tetraacetic acid eNAMPT Extracellular nicotinamide phosphoribosyltransferase ENT Equilibrative nucleoside transporter ER Endoplasmic reticulum ETC Electron transport chain FAD Flavin adenine dinucleotide FADH2 Flavin adenine dinucleotide FBS Fetal bovine serum FOXO1 Forkhead box O 1 G6PD Glucose-6-phosphate dehydrogenase GAPDH Glyceraldehyde phosphate dehydrogenase GDH Glutamate dehydrogenase GIT Gastrointestinal tract Gln Glutamine GPR109a G-protein coupled receptor 109a GPx Glutathione peroxidase GR Glutathione reductase GSH Glutathione HBSS Hanks buffered salt solution HD Helical domain HEK293 Human embryonic kidney cell line HepG2 Human hepatocellular carcinoma cell line HIC1 Hypermethylated in cancer 1 HIF-1α Hypoxia-inducible factor 1 HMGCS2 3-hydroxy-3-methylglutaryl CoA synthase 2 HPLC High performance liquid chromatography HR Homologous recombination I/R Ischemia/reperfusion IDH Isocitrate dehydrogenase IFN-1 Interferon-1 JNK c-Jun N-terminal kinases KFA Kynurenine formamidase KMO Kynurenine 3-hydroxylase KYU Kynureninase LB Luria Bertani LCAD Long-chain acyl coenzyme A dehydrogenase LCFA Long chain fatty acid LC-MS/MS Liquid chromatography tandem mass spectrometry
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Chapter 1 Introduction
LPS Lipopolysaccharide m/z mass to charge ratio MART Mono-ADP-ribosyltransferase MDC1 Mediator of DNA damage checkpoint protein 1 ME Malic enzyme MeOH Methanol MERIT40 Mediator of Rap80 interactions and targeting 40 kd MES Methionine sulfonate MnSOD Manganese superoxide dismutase MPT Mitochondrial permeability transition MRE11 Meiotic recombination 11 MRM Multiple reaction monitoring MRN complex MRE11-RAD50-NSB1 MS Mass spectrometry NA Nicotinic acid NaAD Nicotinic acid adenine dinucleotide NAADP Nicotinic acid adenine dinucleotide phosphate NAD+ Nicotinamide adenine dinucleotide (oxidised form) NADH Nicotinamide adenine dinucleotide (reduced form) NADK Nicotinamide adenine dinucleotide kinase NADSYN Nicotinamide adenine dinucleotide synthetase NAM Nicotinamide NaMN Nicotinic acid mononucleotide NAMPT Nicotinamide phosphoribosyltransferase NAPRT Nicotinic acid phosphoribosyltransferase NaR Nicotinic acid riboside NBS1 Nijmegen breakage syndrome 1 NBTI S-(4-nitrobenzyl)-6-thioinosine NER Nucleotide excision repair NF-κB Nuclear factor kappa B NHEJ Non-homologous end joining NK Natural killer NMN Nicotinamide mononucleotide NMNAT Nicotinamide mononucleotide adenylyltransferase NNT Nicotinamide nucleotide transhydrogenase NOX NADPH oxidase NPP1 Nucleotide pyrophosphatase/phosphodiesterase 1 NPSC Neural/progenitor stem cells NR Nicotinamide riboside NRH Nicotinamide riboside (reduced) NRK Nicotinamide riboside kinase NTPDase Nucleoside tri- and diphosphohydrolase OD Optical density PAR Poly-ADP-ribose
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Chapter 1 Introduction
PARP Poly-(ADP)-ribose polymerase PBMC Peripheral blood mononuclear cells PBS Phosphate buffered saline PD Parkinson's Disease PD-1 Programmed death 1 PDH Pyruvate dehydrogenase PD-L1 Programmed death 1 ligand PGC-1α PPARγ coactivator 1-alpha Pnc1 Nicotinamide deamidase in yeast PncA Nicotinamide deamidase in E. coli PncC Nicotinamide mononucleotide deamidase in E. coli PNKP Polynucleotide kinase 3′-phosphatase PNP Purine nucleoside phosphorylase PPARγ Proliferator-activated receptor gamma PPP Pentose phosphate pathway PRPP 5-phosphoribosyl-1-pyrophosphate PRS Phosphoribosyl synthetase Prx Peroxiredoxins Q1 First quadrupole Q2 Second quadrupole Q3 Third quadrupole QA Quinolinic acid QAPRT Quinolinic acid phosphoribosyltransferase R5P Ribose-5-phosphate rDNA ribosomal DNA ROS Reactive oxygen species RT Retention time SASP Senescence associated secretory phenotype SDH Succinate dehydrogenase SIL Stable isotopically labelled Sir2 Sirtuin 2 in yeast SLC Solute carrier SOD2 Superoxide dismutase SRM Selective reaction monitoring SSB Single-stranded break SUMO Small ubiquitin-like modifier TCA Trichloroacetic acid TD4 Thymine-d4 TDO Tryptophan 2,3-dioxygenase TPCN Two pore calcium channels TR Thioredoxin reductase TRF1 Telomeric repeat factor 1 Trp Tryptophan TRX Thioredoxin
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Chapter 1 Introduction
TRX-S2 Thioredoxin (oxidised) UCP1 Uncoupling protein 1 UPR Unfolded protein response UV Ultraviolet WAT White adipose tissue Wlds Slow Wallerian degeneration XPA Xeroderma pigmentosum group A-complementing protein XPC Xeroderma pigmentosum group C-complementing protein XRCC1 X-ray repair cross-complementing protein 1
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Chapter 1 Introduction
Introduction
1.1 Adding Health to Years
Human lifespan is increasing, but not necessarily in better health. Globally, the greatest causes
of mortality in the elderly are heart disease, stroke and chronic lung disease while the greatest
causes of disability involve sensory impairments, back and neck pain, chronic obstructive pulmonary disease, depressive disorders, falls, diabetes, dementia and osteoarthritis (WHO
2015). The incidence of these and many other age-related diseases and disabilities can be lowered through lifestyle interventions including exercise and moderating nutritional intake, though promoting the widespread uptake of these interventions has met with mixed results.
In the last two decades, the biology of ageing has emerged as an exciting field with the aim of understanding fundamental processes that cause physiological decline with age. Ageing is now recognised as a malleable process, which was first demonstrated through the finding
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Chapter 1 Introduction that a simple reduction in food intake in rats extended overall lifespan (McCay, Crowell, and
Maynard 1935). Since then, work has been undertaken to separate the concepts of biological age versus chronological age. The physical and social aspects of an individuals’ environment can have a significant impact on lifespan, and more importantly, on healthspan—the years of life spent in the absence of disease or disability and maintained in good health (Crimmins
2015).
At a biological level, ageing is characterized by a gradual, lifelong accumulation of molecular and cellular damage that results in a progressive impairment in physiological function, an increased vulnerability to environmental challenges and a growing risk of disease and death (WHO 2015). Importantly, this definition of ageing is also applicable to a number of other disease states, where molecular and cellular damage accumulates with time. On a molecular level, the mechanisms involved in the pathogenesis of individual age related diseases are thought to drive ageing itself, and drawing on these parallels helped define the nine hallmarks of ageing (López-Otín et al. 2013). These hallmarks are genomic instability, telomere attrition, epigenetic alterations, loss of proteostasis, deregulated nutrient sensing, mitochondrial dysfunction, cellular senescence, stem cell exhaustion and altered intercellular communication. The ultimate goal of ageing research is to find interventions that target these hallmarks to increase healthspan during ageing.
Interestingly, the interventions that extend lifespan, through genetic or pharmacological manipulation in experimental models, often result in a broad-spectrum improvement in health and vice versa. The most consistent and robust intervention that demonstrates this is caloric restriction (CR), defined by a reduction in food intake without malnutrition (Speakman and
Mitchell 2011). First demonstrated in rats (McCay, Crowell, and Maynard 1935), CR-induced lifespan extension has been shown across multiple organisms from yeast to humans, revealing that the mechanisms regulating lifespan involve evolutionarily conserved pathways (Anderson
2
Chapter 1 Introduction
et al. 2003; Burger, Buechel, and Kawecki 2010; Civitarese et al. 2007; Colman et al. 2009;
Colman et al. 2014; Kaeberlein et al. 2004; Lee et al. 1999; Lee et al. 2006; Lin et al. 2004;
Mattison et al. 2017; Rogina and Helfand 2004; Witte et al. 2009; Zhang et al. 2012). A lifespan
extension of 2-3-fold was achieved in yeast, fruit flies and worms and up to 50% extension in
median and maximal lifespan in laboratory rodents following a 20-50% caloric reduction. In
addition to lifespan extension, CR delays the onset and progression of various pathological
conditions including cardiomyopathies (Cohen et al. 2017; Niemann et al. 2010), diabetes
(Baumeier et al. 2015; Hammer et al. 2008; Teeuwisse et al. 2012), cancer (Cadoni et al. 2017;
Lanza-Jacoby et al. 2013) and neurodegeneration (Gräff et al. 2013).
Due to the challenges of implementing CR in daily life, considerable effort has been invested to find interventions that mimic the benefits of CR. For example, the anti-diabetic drug metformin has demonstrated a longevity benefit independent of its effect in glycaemic control (Anisimov et al. 2011a; Cabreiro et al. 2013; Martin-Montalvo et al. 2013), with various improved aspects of health (Haak et al. 2012; Martin-Montalvo et al. 2013; Morin-Papunen et
al. 2012) including the risk of certain cancers (Mohammed et al. 2013; Shankaraiah et al. 2019), kidney disease (Neven et al. 2018), neurodegeneration (Lu et al. 2016) and cardiac dysfunction
(Pandey et al. 2019), among many others. In addition to metformin, other CR-mimetics that show similar benefits to healthspan and lifespan include resveratrol (Rascón et al. 2012), rapamycin (Anisimov et al. 2011b; Bitto et al. 2016; Harrison et al. 2009), spermidine
(Eisenberg et al. 2016; Eisenberg et al. 2009), quercetin (Alugoju et al. 2018), and intermittent fasting (Catterson et al. 2018; Honjoh et al. 2009; Mitchell et al. 2019), among others.
There are four key proteins that have been identified as mediating the benefits of CR.
These include the insulin/insulin-like growth factor 1 (insulin/IGF-1), adenosine monophosphate activated protein kinase (AMPK), mechanistic target of rapamycin (mTOR) and the sirtuins (SIRTs) (Henrique Mazucanti et al. 2015; Pan and Finkel 2017). Briefly,
3
Chapter 1 Introduction
insulin/IGF-1 is part of the insulin/IGF-1 signalling (IIS) pathway, a highly conserved nutrient
sensing pathway regulated by hormones such as insulin and growth hormone (GH) which
regulates IGF-1 levels. One of the mechanisms to explain CR-induced lifespan extension is
thought to be through reducing plasma and tissue levels of insulin and IGF-1 (Li et al. 2011;
Zaarur et al. 2019; Zarse et al. 2012). While this has sparked controversy in the past, with some
arguing that lifespan extension through impaired IIS signalling is independent of CR-mediated lifespan extension (Houthoofd et al. 2003; Min et al. 2008), recent studies continue to show that downregulation of insulin/IGF-1 levels are still relevant to the beneficial effects of CR in lifespan extension (Zaarur et al. 2019; Zarse et al. 2012). The nutrient and energy sensor AMPK is activated by metabolic stressors that decrease energy availability, such as CR. Once activated, it promotes compensatory utilisation of other energy substrates for ATP generation, increasing the beta oxidation of fatty acids, and activates other proteins such as the sirtuins and mTOR and the IIS pathway to mediate its protective effects (Greer et al. 2007; Hardie, Ross, and Hawley 2012; Palacios et al. 2009). mTOR is also an important cellular nutrient and energy sensor activated in response to an abundance of nutrients (amino acids) or stimulated by growth factors to promote growth and proliferation by enhancing anabolic processes such as translation and ribosomal biogenesis, while inhibiting catabolic processes such as autophagy. The inhibition of mTOR, through the clinically used immunosuppressive small molecule rapamycin, extends lifespan (Anisimov et al. 2010) and the benefits of CR are also considered to be mediated in part by the downregulation of mTOR (Dogan et al. 2011; Yang et al. 2014).
The sirtuins are a family of seven histone deacylases whose activity is dependent on the consumption of nicotinamide adenine dinucleotide (NAD+) (Frye 1999; Imai et al. 2000a;
Landry et al. 2000). They play a major role in response to injury and stress and regulate almost
all cellular functions such as energy metabolism, inflammation and neuronal function and play
a major role in mediating lifespan extension (Houtkooper, Pirinen, and Auwerx 2012; Howitz
4
Chapter 1 Introduction
et al. 2003; Jęśko et al. 2017; Vachharajani et al. 2016). In recent years, the redox and enzyme
cofactor NAD+ has emerged as an important determinant of late-life health and lifespan (Fang
et al. 2017; Johnson and Imai 2018; Zhang et al. 2016a). NAD+ levels decline with age or in
various pathological conditions, and strategies that elevate NAD+ levels have created
promising new avenues to combat age-related diseases (Aman et al. 2018; Rajman, Chwalek, and Sinclair 2018; Yoshino, Baur, and Imai 2018).
1.2 The Biological Significance of NAD+
NAD+ is a pyridine nucleotide first discovered in 1906 as a cofactor for enhancing alcohol
fermentation in yeast (Harden and Young 1906a, 1906b). Shortly after, NAD+ was successfully
isolated from yeast extracts by Hans von Euler-Chelpin in 1936 who characterised its chemical structure as a pyridine nucleotide made up of two mononucleotides – adenosine monophosphate (AMP) and nicotinamide mononucleotide (NMN) (Figure 1.1, below).
Figure 1.1 Chemical structure of NAD+ Nicotinamide adenine dinucleotide (NAD+) is made up of two mononucleotides, adenosine monophosphate (AMP) and nicotinamide mononucleotide (NMN) (Schlenk and Euler 1936).
5
Chapter 1 Introduction
In the same year, Otto Warburg discovered the function of NAD+, as a molecule capable
of transferring electrons between compounds in metabolic reactions. Its phosphorylated form,
NADP+, was also discovered (Warburg and Christian 1936) and this compound has important functions as a redox cofactor and in anabolic reactions, especially in its reduced form, NADPH
(Agledal, Niere, and Ziegler 2010). In 1950, Arthur Kornberg was the first to show that NAD+
was synthesised from the condensation of ATP and NMN via the enzyme nicotinamide
mononucleotide adenylyltransferase (NMNAT) (Kornberg 1950). Since these discoveries, the
diverse roles of NAD+ continue to expand and emerge as an molecule of central importance in
molecular and cell biology.
Today, NAD+ is considered one of the most fundamental components in life involved
in over 500 enzymatic reactions in the body (Ansari and Raghava 2010). NAD+ levels decline
with age (Braidy et al. 2011b; Clement et al. 2019; Gomes et al. 2013; Massudi et al. 2012a;
Mouchiroud et al. 2013; Stein and Imai 2014) and are also implicated in pathological conditions such as diabetes (Trammell et al. 2016c; Yoshino et al. 2011), obesity (Cantó et al. 2012), non- alcoholic fatty liver disease (Zhou et al. 2016), and neurodegeneration (Hou et al. 2018; Lu et al. 2014a). NAD+ levels can be increased either through genetic manipulation or through
pharmacological supplementation with NAD+ precursors, resulting in benefits that are
mediated by its critical role as an essential cofactor used in oxidation-reduction (redox) reactions, and secondly, as a crucial substrate and modulator of NAD+-consuming enzymes
that regulate biological processes from energy metabolism to DNA repair and cell survival
(Braidy et al. 2019; Kulkarni and Brookes 2019; Xiao et al. 2018) (Figure 1.2, below).
Interestingly, NAD+ levels oscillate according to the circadian clock, an autonomous molecular
programme that drives the expression of thousands of genes around a 24-hour cycle. This
circadian cycle regulates the expression of nicotinamide phosphoribosyltransferase (NAMPT),
6
Chapter 1 Introduction the rate-limiting enzyme involved in NAD+ biosynthesis (Ramsey et al. 2009) (Figure 1.2, below).
Figure 1.2 The biological roles of NAD+ NAD+ is involved in over 500 enzymatic reactions in the body. NAD+ is an important cofactor in (1) oxidation-reduction (redox) reactions involved in key metabolic processes such as (2) glycolysis, reactions in the tricarboxylic acid (TCA) cycle and oxidative phosphorylation (OXPHOS). NAD+ is also an important substrate for NAD+-consuming enzymes such as (3) poly-ADP-ribose polymerases (PARPs) involved in DNA damage repair and (4) the sirtuins (SIRTs), histone deacetylases that regulates gene expression and respond to cellular stress. NAD+ is also degraded by glycohydrolases (NADases) such as (5) cluster of differentiation number 38 (CD38) and its homologue, CD157, that play a role in calcium (Ca2+) mobilization and the immune response. (6) NAD+ is also degraded by a protein called sterile alpha and Toll/interleukin-1 receptor motif-containing 1 (SARM1) which is implicated in axonal degeneration. (7) NAD+ levels also oscillate around a 24-hour cycle governed by the circadian clock, an autonomous molecular programme that drives the expression of thousands of genes around a 24-hour cycle.
7
Chapter 1 Introduction
1.2.1 NAD+ as a redox cofactor
NAD+ plays a fundamental role as a redox cofactor to donate or accept electrons, rapidly
shuttling between its oxidised (NAD+) and reduced (NADH) forms, in its dephosphorylated
state as NAD+ and NADH, or in its phosphorylated state as NADP+ and NADPH (Figure 1.3,
below).
1.2.1.1 NAD+ and NADH roles and functions
Redox reactions are primarily carried out by dehydrogenase, hydroxylase and reductase
enzymes in the cell and involves the transfer of electrons between metabolites (Monti et al.
2011). NAD+ severs as an electron acceptor, which is reduced to form NADH. Similarly,
NADH serves as an electron donor which is oxidised to regenerate NAD+. It is important to
recognise that redox reactions between NAD+ and NADH simply involve the exchange of
electrons in a non-destructive way. In contrast, the role of NAD+ as a substrate, discussed in
the next section, involves the hydrolysis of NAD+, requires its resynthesis from precursors in
a process that also consumes energy in the form of ATP.
NAD+ is reduced in major energy production pathways such as glycolysis (via glyceraldehyde phosphate dehydrogenase; GAPDH) (Harris 1972), pyruvate oxidation to acetyl-CoA (via pyruvate dehydrogenase; PDH) (Wieland 1983), the tricarboxylic acid (TCA) cycle (via α-ketoglutarate, isocitrate and malate dehydrogenases) (Cupp and McAlister-Henn
1992; Hägele, Neeff, and Mecke 1978; Pettit et al. 1973; Seelig and Colman 1978) and fatty acid oxidation (via 3-hydroxyacyl-CoA dehydrogenase) (Uchida et al. 1992). Another reduced cofactor, flavin adenine dinucleotide (FADH2) is generated alongside NADH in the TCA cycle
by reducing the oxidised form, flavin adenine dinucleotide (FAD) via succinate dehydrogenase
(Kita et al. 1989). NADH and FADH2 are both important cofactors in energy metabolism as
they donate their electrons to complex I and II of the electron transport chain (ETC),
8
Chapter 1 Introduction
respectively, to generate adenosine triphosphate (ATP) during oxidative phosphorylation
(OXPHOS) in the mitochondria (Papa et al. 2012). NADH made in the cytoplasm is shuttled
into the mitochondria via two separate shuttle systems: the malate-aspartate shuttle which is
reversible and relies on high cytosolic NADH levels, or the glycerol phosphate shuttle which
is irreversible and can act in low cytosolic NADH levels (Luo, Li, and Yan 2015). Delivery of
electrons to the ETC is an important step for regenerating NAD+ to maintain intracellular NAD+
levels and the NAD+/NADH ratio, which has been recorded at approximately 700 in the cytoplasm and 7-8 in the mitochondria (Stubbs, Veech, and Krebs 1972). An imbalance in the
NAD+/NADH ratio, usually from an excess of NADH can lead to increased oxidative stress,
and has been implicated in various pathological conditions such as diabetes (Wu, Jin, and Yan
2017; Wu et al. 2017), acute kidney injury (Nechifor and Dinu 2011), and heart failure (Lee et
al. 2016a).
1.2.1.2 NADP+ and NADPH roles and functions
An estimated 10% of the total NAD+ and NADH pool exists in their phosphorylated forms,
NADP+ and NADPH. The non-phosphorylated cofactors NAD+ and NADH primarily participate in catabolic reactions while their phosphorylated forms, NADP+ and NADPH are
mainly required in anabolic reactions and cellular oxidative stress defence.
NADP+
The main source of NADP+ is from the phosphorylation of NAD+ via ATP-dependent NAD
kinase (NADK) in the cytoplasm (Lerner et al. 2001) or mitochondria (Ohashi, Kawai, and
Murata 2012). As NADP+ is a crucial substrate to generate NADPH, which in turn is a key
regenerator of antioxidants in the cell, activity of NADK is also crucial. The downregulation
of NADK is associated with increased sensitivity to reactive oxygen species (ROS) such as
hydrogen peroxide (H2O2) (Pollak, Niere, and Ziegler 2007) and can lead to hepatic steatosis
9
Chapter 1 Introduction in mice (Koh et al. 2004; Zhang et al. 2018b). Overexpression of NADK increases NADPH production 4-5-fold, and provides increased protection against oxidative stress (Pollak, Niere, and Ziegler 2007).
Under normoglycemia, glycolysis is the main pathway for glucose metabolism but a small amount filters into the polyol pathway where glucose is converted into sorbitol via the rate-limiting enzyme aldose reductase (AR), a step which oxidises NADPH to form NADP+
(Wermuth et al. 1982). The polyol pathway is also a source of NADH in the next immediate step where sorbitol is converted into fructose via sorbitol dehydrogenase, reducing NAD+ in the process. Under hyperglycaemic conditions, there is increased glucose flux through the polyol pathway which not only depletes NADPH but also generates NADH. As NADPH is an important cofactor in regenerating cellular antioxidants, increased NADPH oxidation through the polyol pathway will lead to impaired antioxidant regeneration and an accumulation of ROS.
Further, excess NADH will also contribute to more oxidative damage as NADH oxidation through complex I leads to increased leakage of electrons from the ETC, which react with oxygen to create superoxide anions and H2O2, which are sources of ROS stress. Oxidative stress induced by overactivation of the polyol pathway is implicated in pathological conditions including diabetes (Snow et al. 2015; Son et al. 2012; Tang et al. 2014). Consistent with this, treatment with AR inhibitors have prevented or delayed the progression of various disease conditions and diabetic complications (Hotta et al. 2012; Kawai et al. 2010; Qiu et al. 2011;
Sonowal et al. 2017; Srivastava et al. 2011).
NADP+ is also the precursor for the synthesis of nicotinic acid adenine dinucleotide phosphate (NAADP) (Chini and Dousa 1995), a vital second messenger for intracellular calcium mobilisation (Calcraft et al. 2009; Steen, Kirchberger, and Guse 2007). The release of calcium from intracellular stores is important for biological processes including lysosomal
10
Chapter 1 Introduction
function (Lu et al. 2013a; Zong et al. 2009), autophagy (Lu et al. 2013b; Pereira et al. 2011)
and activation of the mTOR pathway when cellular energy levels are low (Cang et al. 2013).
NADPH
The majority of NADPH is generated by the rate limiting enzyme glucose-6-phosphate
dehydrogenase (G6PD) in the oxidative pentose phosphate pathway (PPP) (Kirkman 1962;
Yoshida 1966). G6PD catalyses the rate-limiting step of the PPP, where its substrate glucose-
6-phosphate (G6P) is converted into 6-phosphogluconate (6-PG), reducing NADP+ to NADPH
in the process. The next immediate step also generates NADPH, where 6-phosphogluconate
dehydrogenase (6PGD) (Pearse and Rosemeyer 1974, 1975) catalyses the conversion of 6-PG
to form ribose-5-phosphate (R5P), an important building block for nucleotide biosynthesis
(Blakley and Vitols 1968). In addition to the PPP, NADPH can also be synthesised via
isocitrate dehydrogenase (IDH) and malic enzyme (ME) which have cytosolic (IDH1, ME1)
(Geisbrecht and Gould 1999; Loeber et al. 1994) and mitochondrial (IDH2, ME3) isoforms
(Loeber et al. 1991; Luo, Shan, and Wu 1996). IDH1/2 catalyses the conversion of isocitrate
to α-ketoglutarate while ME1/3 catalyses the conversion of malate to pyruvate, both reducing
NADP+ to generate NADPH.
Other NADP+-dependent mitochondrial enzymes such as glutamate dehydrogenase
(GDH) and nicotinamide nucleotide transhydrogenase (NNT) also contribute to the NADPH
pool in mitochondria. GDH is commonly upregulated in many cancers (Jin et al. 2015; Spinelli
et al. 2017), promoting tumour growth through increased antioxidant activation and redox
scavenging. Inhibition of GDH activity has been a promising new development in anti-cancer therapeutics (Hou et al. 2019). NNT is a nuclear-encoded protein located in the inner mitochondrial membrane (Arkblad et al. 2002), where it serves to maintain redox homeostasis in the mitochondria in an NADPH-manner, and its deficiency slows tumour growth and reduces
11
Chapter 1 Introduction
tumorigenicity in cells (Ho et al. 2017), also making them prime therapeutic targets for treating
cancer (Chortis et al. 2018; Li et al. 2018).
NADPH acts as an essential cofactor in thioredoxin and glutaredoxin systems to
manage ROS-induced oxidative stress (Jacquot and Zaffagnini 2019). Glutathione reductase
(GR) (Carlberg et al. 1975) and thioredoxin reductase (TR) (Luthman et al. 1982) use NADPH
as a reducing factor to regenerate glutathione (GSH) and thioredoxin (TRX) from their oxidised
forms, GSSH and oxidised thioredoxin (TRX-S2), respectively. Glutathione peroxidases (GPx)
and peroxiredoxins (Prx) then assist in the removal of ROS in the cell by transferring electrons
from GSH and TRX onto oxidative species such as H2O2 forming water. The major source of
ROS in the cell comes from the leakage of electrons out of ETC into the mitochondrial matrix
during oxidative phosphorylation (Vinogradov and Grivennikova 2016). These electrons are
accepted by molecular oxygen, as it is highly electronegative, producing ROS such as
superoxide or H2O2. The removal of these cytotoxic species requires rapid detoxification by
NADPH-GSH-Gpx and NADPH-TRX-Prx systems where an impairment in their coordinated activity is associated with increased oxidative stress and cellular damage (Esposito et al. 2000;
Jin et al. 2011; Oelze et al. 2014).
Another group of enzymes called NADPH oxidases (NOXs) catalyse the production of
- superoxide free radicals (O2 ) by using NADPH as a reducing agent to transfer electrons to
+ + - oxygen in the following reaction: NADPH + 2O2 NADP + H + 2O2 . Under basal
conditions, NOXs are considered inactive, but become activated in response to infection by
pathogens. NOXs are typically embedded in the plasma membrane of phagocytes such as
neutrophils, eosinophils, monocytes and macrophages. In the presence of a pathogen, they
release of large amounts of ROS into the phagosome in an important immune response process
known as a respiratory burst (El‐Benna et al. 2016; Thomas 2017). Unlike other instances
where the generation of ROS and free radicals is highly detrimental to cells, respiratory bursts
12
Chapter 1 Introduction act to destroy extracellular pathogens such as bacteria or fungi ingested by the phagosome, serving as a protective mechanism for the cell. NOXs can be highly detrimental as they generate large amounts of superoxides and their overaction can lead to increased oxidative stress and cellular damage (Almenara et al. 2015; Babu et al. 2015; Stanicka et al. 2015). This has been implicated in various age-related conditions involving the heart (Barman et al. 2014; Di Marco et al. 2014; Wang et al. 2014a), brain (Bruce-Keller et al. 2010; Cooney, Bermudez-Sabogal, and Byrnes 2013; Pal et al. 2014; Shen et al. 2016), liver (Choi et al. 2014), kidney (Alhasson et al. 2017; Djamali et al. 2009; Gao et al. 2016) and loss of bone (Chen et al. 2011a).
Modulating NADPH levels through the inhibition of NOX isoforms attenuates oxidative damage and pathological progression (Cha et al. 2017; Chen et al. 2011a; Di Marco et al. 2014;
Kwon et al. 2017; Lu et al. 2014b; Qin et al. 2017).
Overall, NAD in its oxidised, reduced and phosphorylated forms are involved in diverse redox roles and functions accounting for over 400 enzymatic reactions in the body, highlighting its role as a central regulator of energy metabolism and redox homeostasis in cells.
13
Chapter 1 Introduction
Figure 1.3 Functions of NAD(P)(H) as a redox cofactor. The oxidised form of nicotinamide adenine dinucleotide (NAD+) participates in oxidation- reduction (redox) reactions in processes such as glycolysis, β-oxidation, fatty acid oxidation and reactions in the TCA cycle to produce NADH. NADH is a major carrier of electrons and donates its electrons to the electron transport chain (ETC) during oxidative phosphorylation in the mitochondria, regenerating NAD+ in the process. NAD+ can be phosphorylated to form NADP+ via NAD kinase (NADK) which requires energy in the form of adenosine triphosphate (ATP) and forms adenosine diphosphate (ADP) as a by-product. NADK also catalyses the phosphorylation of NADPH from NADH in an ATP-dependent manner. The generation of NADP+ is important for generating its reduced form NADPH, which plays a key role in regeneration of antioxidants such as glutathione. The main enzymes that generate NADPH from NADP+ are dehydrogenases in the Pentose phosphate pathway (PPP). In the reverse reaction, NADPH can be oxidised to NADP+ by NADPH oxidases and NADPH- dependent reductases. Recently, the enzyme nocturnin, which regulates metabolism under the control of circadian clock was shown to catalyse the dephosphorylation of NADPH and NADP+ to generate NADH and NAD+ respectively, and is conserved from fruit flies to humans (Estrella et al. 2019).
14
Chapter 1 Introduction
1.2.2 NAD+ as a substrate
In addition to the role of NAD+ as a redox cofactor, in the last two decades it has emerged as a
key signalling molecule and substrate capable of modulating the activity of important NAD+-
consuming enzymes. These include 1) poly-ADP-ribose polymerases (PARPs), 2) the sirtuins,
3) NAD+ glycohydrolases (CD38/CD157) and 4) sterile alpha and Toll/interleukin-1 receptor
motif-containing 1 (SARM1) (Figure 1.2, above). All cleave NAD+ at the nicotinamide-ribose bond to form free nicotinamide (NAM) and an ADP-ribose (ADPR) moiety. NAM can be recycled back into the salvage pathway to synthesise more NAD+ molecules while ADPR is used by these enzymes to carry out their respective biological functions.
1.2.2.1 NAD+ and PARPs
In 1963, nearly three decades after discovering the role of NAD+ as a redox cofactor, NAD+
was discovered as the immediate substrate to a family of enzymes called poly-ADP-ribose
polymerases (PARPs) (Chambon, Weill, and Mandel 1963; Kraus 2020) or mono-ADP-ribose transferases (MARTs), when only one ADP-ribose molecule is transferred to proteins
(Bütepage et al. 2015; Ueda and Hayaishi 1985). PARPs are responsible for the addition of polymers of adenosine diphosphate (ADP)-ribose to proteins (PARylation) (Chambon et al.
1963). Specifically, they target the amino acid residues on proteins such as arginine,
asparagine, aspartate, cysteine, glutamate, lysine, and serine (Leidecker et al. 2016; Manning
et al. 1984; Martello et al. 2016; Moss et al. 1979; Ogata, Ueda, and Hayaishi 1980; West et
al. 1985; Zhang et al. 2013). PARylated proteins act as a signal and docking site for protein
interactions with proteins involved in processes such as DNA repair.
In humans, there are 17 members of the PARP family, each with unique function and
location. Briefly, the members of the PARP family that are involved in the addition of polymers
of ADP-ribose (PARylation) are PARP1, PARP2, PARP5a (also known as tankyrase 1) and
PARP5b (also known as tankyrase 2) while most other PARP members (PARP3, 4, 6-16)
15
Chapter 1 Introduction
remain as MARTs (Cohen 2020; Fehr et al. 2020; Vyas et al. 2014). PARPs are located in the
nucleus (PARP1-3, 6-10, 15), nucleolus (PARP1 and 2), nuclear envelope (PARP11),
cytoplasm (PARP4-6, 8-10, 14), Golgi (PARP12), and endoplasmic reticulum (PARP16)
(Bindesbøll et al. 2016; Carter-O’Connell et al. 2016; Carter-O’Connell et al. 2018; Catara et
al. 2017; Di Paola et al. 2012; Gomez et al. 2018; Jwa and Chang 2012; Vyas et al. 2013),
where their role is discussed in following sections.
PARPs in DNA damage repair
PARPs involved in DNA repair (mainly PARP1) are activated in response to DNA damage
derived from endogenous sources such as the production of ROS from oxidative
phosphorylation in the mitochondria, or from exposure to environmental toxins such as X-rays,
ultraviolet (UV) light, radiation, carcinogens and cigarette smoke (Lindahl 1993; Tubbs and
Nussenzweig 2017). PARP1 is the founding member of the PARP family, which is best known
for its role in DNA damage repair (DDR) (El‐Khamisy et al. 2003; Haince et al. 2008; Le Page
et al. 2003; Mortusewicz et al. 2007).
PARP1 consists of three major domains that mediate its catalytic activity. The first is
the DNA binding domain, which consists of an N-terminal three-zinc finger domain (Zn1, Zn2,
and Zn3) and the WGR domain, named after the most conserved amino acid sequence
(tryptophan–glycine–arginine) (Langelier et al. 2012; Langelier et al. 2018). This domain
allows PARP1 to bind to cruciform DNA, which are inverted repeats of nucleotide sequences
(Brázda et al. 2011; Chasovskikh et al. 2005; Potaman et al. 2005) as well as single stranded
DNA breaks (SSBs) (Heale et al. 2006; Le Page et al. 2003; Pines et al. 2012) and double stranded DNA breaks (DSBs) (Audebert, Salles, and Calsou 2004; Beck et al. 2014b; Wang et al. 2006a). The second domain is a central auto-modification domain which includes a BRCA1
C-terminus (BRCT) domain that allows the recruitment of downstream DDR components to
16
Chapter 1 Introduction
the DNA damage site and for protein-protein interactions to occur between PARP1 and DDR
machinery (Ahel et al. 2009; El‐Khamisy et al. 2003; Haince et al. 2008; Langelier et al. 2012;
Li and Yu 2013; Masson et al. 1998). This auto-modification domain also allows PARPs such
as PARP1 and PARP2 to auto-PARylate themselves, which helps them dissociate from DNA
when damage is repaired, demonstrating a self-regulated negative feedback mechanism (Liu et
al. 2017b; Masaoka et al. 2012; Muthurajan et al. 2014; Satoh and Lindahl 1992; Sukhanova
et al. 2016).
The final structural domain in PARPs is a C-terminal catalytic domain which consists
of two subdomains: a helical domain (HD) and an ADP-ribosyltransferase (ART) catalytic
domain (Dawicki-McKenna et al. 2015; Langelier et al. 2018). When not bound to DNA, the
HD acts as an autoinhibitory domain by sitting in a conformation that prevents NAD+ from binding to the ART catalytic domain (Dawicki-McKenna et al. 2015). When bound to DNA
breaks, the HD again changes conformation such that the autoinhibitory function is lost,
allowing NAD+ binding which activates the catalytic activity of PARP1 (Dawicki-McKenna et
al. 2015). The use of clinical PARP inhibitors to treat diseases such as cancer target and bind
to the ART catalytic site, preventing its release from DNA and trapping PARP1 at the DNA
damage site to induce cell death (Lord and Ashworth 2017; Murai et al. 2014; Murai et al.
2012).
PARP1-mediated SSB repair can occur via base excision repair (BER) (Dantzer et al.
1999) or nucleotide excision repair (NER) (King et al. 2012). BER involves the removal of a
single or short length of damaged or misplaced DNA base(s) (A-C-T-G). The main protein
interacting with PARP1 to perform BER is X-ray repair cross-complementing protein 1
(XRCC1) which recruits other important DNA intermediates including DNA polymerase β
(Dianova et al. 2004; Kubota et al. 1996; Sobol et al. 1996), DNA ligase III (Cappelli et al.
1997), bifunctional polynucleotide kinase 3′-phosphatase (PNKP) (Jilani et al. 1999; Mandal
17
Chapter 1 Introduction et al. 2012) and DNA glycohydrolases (Krokan, Standal, and Slupphaug 1997; Mandal et al.
2012). XRCCI and DNA ligase III are also indispensable during NER which seals breaks in the DNA following repair (Moser et al. 2007). NER removes larger sites of DNA damage called oligonucleotide fragments. These fragments are initially recognised by proteins xeroderma pigmentosum group C-complementing protein (XPC) (Maltseva et al. 2015; Robu et al. 2013;
Robu et al. 2017) and xeroderma pigmentosum group A-complementing protein (XPA)
(Fischer et al. 2014; King et al. 2012) which recruit other proteins to form a large multiprotein complex, called DNA damage-binding protein 2 (DDB2) complex (Purohit et al. 2016). PARP1 interacts with DDB2 at DNA damage sites, stimulating its PARylation activity to recruit other chromatin remodelling factors such as amplified in liver cancer protein 1 (ALC1) which stimulates the DNA repair process (Pines et al. 2012). PARP2 is also a key player in BER by binding to PARP1, which assists in the recruitment of core DDR proteins, XRCC1, DNA polymerase β and DNA ligase III for efficient BER (Hanzlikova et al. 2017; Schreiber et al.
2002).
DSBs occur after exposure to DNA damaging agents such as UV radiation (Mehta and
Haber 2014; Rothkamm et al. 2003). DSBs are repaired either via homologous recombination
(HR) (Schultz et al. 2003) or non-homologous end joining (NHEJ) (Audebert, Salles, and
Calsou 2008; Wang et al. 2006a). HR repairs DNA during the S and G2 phase of the cell cycle
(Mao et al. 2008) using the sister chromatid as a template for repair and replacing the damaged
DNA with an identical nucleotide sequence with a high level of fidelity. In contrast, NHEJ is most efficient in the G1 phase (Mao et al. 2008; Moore and Haber 1996), and is an error prone repair pathway as it does not use a complementary template but simply ligates the break ends of DNA.
In both HR and NHEJ, PARP1 is involved in the recruitment of DNA damage sensors meiotic recombination 11 (MRE11) and Nijmegen breakage syndrome protein 1 (NBS1 or
18
Chapter 1 Introduction
nibrin), which rapidly accumulates at DNA damage sites following DSBs (Haince et al. 2008).
PARP1 is also involved in the early recruitment of breast cancer type 1 susceptibility protein
(BRCA1), which recruits other DDR proteins such as RAD51 to the damage site during HR
(Chen et al. 1999; Zhao et al. 2017). When the sister chromatid is not available for HR, DSBs
are repaired via NHEJ which has two different pathways: the classical (cNHEJ) or alternative
(aNHEJ) pathway. cNHEJ is dependent on the presence of DNA repair dimer Ku70-Ku80 that
recruit the DNA-dependent protein kinase catalytic subunit (DNA-PKcs), which when
PARylated recruits XRCC4/DNA ligase IV complex for DNA ligation (Ruscetti et al. 1998;
Spagnolo et al. 2012). In the absence of Ku70-Ku80, the aNHEJ pathway is activated and relies on PARP1-mediated recruitment of MRE11 in the MRE11-RAD50-NSB1 (MRN) complex for
DNA end ligation (Cheng et al. 2011; Fattah et al. 2010; Mansour, Rhein, and Dahm-Daphi
2010; Wang et al. 2006a). PARP3, a MART, helps to recruit and stabilise Ku80 at DNA damage sites to encourage DSB repair by stimulating the cNHEJ pathway (Beck et al. 2014a).
Two others MARTs, PARP10 and PARP14 play roles in alleviating replication stress by coordinating a restart to stalled replication forks that occur as a result of breaks in DNA during replication, or by interacting with RAD51 to promote DNA ligation via HR (Nicolae et al. 2015; Nicolae et al. 2014; Shahrour et al. 2016). While the PARPs responsible for DNA repair processes are essential for the survival of normal cells, they may be undesirable in the context of cancer by promoting cell proliferation and tumorigenesis (Schleicher et al. 2018).
The tankyrases PARP5a and PARP5b indirectly play a role in DDR by binding to DDR proteins
such as mediator of DNA damage checkpoint protein 1 (MDC1) (Goldberg et al. 2003; Lou et
al. 2006; Lou et al. 2003; Stewart et al. 2003; Stucki et al. 2005), mediator of Rap80 interactions
and targeting 40 kd (MERIT40) (Feng, Huang, and Chen 2009; Shao et al. 2009), or telomeric
repeat factor 1 (TRF1) (Smith et al. 1998; Yang et al. 2017b) that are collectively responsible
19
Chapter 1 Introduction for the recruitment and stabilisation of other DDR proteins to the damage site for efficient DNA repair.
PARP1 is the main member involved in DNA damage repair, however, as mentioned above PARP2, 3, 5a, 5b, 9, 10 and 14 also play a role in DNA repair processes (Boehler et al.
2011; Cook et al. 2002; Dregalla et al. 2010; Haince et al. 2008; Nicolae et al. 2014; Schreiber et al. 2002; Yang et al. 2017a). In addition to repairing DNA, PARPs are involved in several other important biological processes. PARP5a and 5b are important in the regulation of telomere length (Cook et al. 2002) and the Wnt signalling, a pathway commonly implicated in cancers (Bao et al. 2012; Huang et al. 2009; Wu et al. 2016). PARP6 is involved in dendritic development in the hippocampus (Huang et al. 2016) and acts as a tumour suppressor in the development of colorectal cancer (Qi et al. 2016; Tuncel et al. 2012). PARP7, 10, 12, 13 and
14 are involved in RNA processing including rDNA synthesis, ribosomal biogenesis, mRNA transcription and protein translation (Bock, Todorova, and Chang 2015). PARP9, 10 and 14 regulate the transcription of genes involved in macrophage activation and inflammation (Iwata et al. 2015; Iwata et al. 2016; Iwata et al. 2014) while PARP1, 5a, 7, 9-14 all play a role in the host response to viral infections (Fehr et al. 2020; Kuny and Sullivan 2016). PARP9 was also found to be overexpressed in human breast cancer and promoted cancer cell migration (Tang et al. 2018). In contrast, PARP10 seems to play a protective role by controlling cell proliferation (Chou, Chou, and Lee 2006; Yu et al. 2005), suppressing tumour metastasis (Zhao et al. 2018) and is beneficial to mitochondrial function when downregulated (Marton et al.
2018). It has been revealed that PARP11 is important for the proper development of sperm
(Meyer-Ficca et al. 2015), organisation of the nuclear envelope (Kirby et al. 2018) and is a potent regulator of type 1 interferon (IFN-I) anti-viral efficiency (Guo et al. 2019). Though the functions of PARP15 remain unclear, a genetic association of PARP15 polymorphisms have been found in patients with acute myeloid leukemia (Lee et al. 2016b). Finally, PARP16 is
20
Chapter 1 Introduction
essential in the regulation of the unfolded protein response (UPR) (Jwa and Chang 2012) and
its localisation to the endoplasmic reticulum (ER), has also been targeted to enhance ER stress-
induced cancer cell apoptosis (Di Paola et al. 2012; Wang et al. 2017a).
The ADP-ribosylating functions of PARPs are highly dependent on hydrolysis of NAD+
molecules and hyperactivation of PARPs can lead to a significant decrease in intracellular
NAD+ (Alano et al. 2010; Du et al. 2003; Pillai et al. 2005; Satchell et al. 2003; Wang et al.
2013), as discussed in later sections. PARP-induced depletion of intracellular NAD+ has been associated with increased cell death (Du et al. 2003; Ha and Snyder 1999; Pillai et al. 2005;
Ying, Garnier, and Swanson 2003; Zhang et al. 2014a) and use of PARP inhibitors protected against PARP-induced NAD+ depletion and cell death (Martín-Guerrero et al. 2017).
1.2.2.2 NAD+ and the sirtuins
The sirtuins are another family of enzymes which regulate cellular stress response, metabolism
and promote healthspan and lifespan extension, consuming NAD+ in the process (Frye 1999;
Imai et al. 2000b; Kaeberlein, McVey, and Guarente 1999; Landry et al. 2000). Silent
information regulator 2 (Sir2) was the founding member of this family, first discovered in the
budding yeast Saccharomyces cerevisiae as a transcriptional repressor of the mating type loci
HML and HMR (Klar and Fogel 1979; Rine et al. 1979; Shore, Squire, and Nasmyth 1984).
Further studies into yeast Sir2 revealed that it prevented the toxic accumulation of senescence-
related ribosomal DNA (rDNA) circles by suppressing recombination at the rDNA locus
(Sinclair and Guarente 1997), with later work showing that it could protect telomeres by
maintaining their length and attenuating age-related telomere shortening (Moretti et al. 1994;
Strahl-Bolsinger et al. 1997; Xu et al. 2007).
In mammals, there are seven known sirtuins (SIRT1-7), of which SIRT1 is the closest mammalian homologue to yeast Sir2 (Brachmann et al. 1995). In 1999, Frye and colleagues
21
Chapter 1 Introduction
discovered that mammalian sirtuins are dependent on NAD+ (Frye 1999) and shortly after, their role in histone deacetylation was also revealed (Imai et al. 2000b; Landry et al. 2000).
The sirtuins are well known for their role in histone deacetylation and play a major role as epigenetic regulators of gene expression. In addition to removing acetyl groups, they also remove other acyl groups including malonyl, succinyl, crotonyl and propionyl from various protein substrates, most notably histones, consuming NAD+ in a reaction that yields NAM and
2’-O-acetyl-ADPR (Imai et al. 2000b; Tanner et al. 2000). NAM serves as a substrate to the
salvage pathway enzyme NAMPT, where it is rate-limiting for NAD+ biosynthesis (Revollo,
Grimm, and Imai 2004), while ADPR accepts the acyl groups that are removed by the sirtuins to form 2’-O-acetyl-ADPR (Tanner et al. 2000). Importantly, high concentrations of NAM inhibit the deacetylase activity of the sirtuins via a negative feedback loop (Bitterman et al.
2002), highlighting the importance of the salvage pathway in using NAM not just as a substrate
to restore NAD+ levels, but to potentially avoid the inhibition of sirtuin activity.
Importantly, the beneficial effects of this activity is only achieved if NAD+ is readily
bioavailable for their consumption, and a decline or depletion in intracellular NAD+ levels is
concurrent with decreased sirtuin activity and downregulation of the processes they control
(Braidy et al. 2011b; Cantó et al. 2009; Fang et al. 2014; Furukawa et al. 2007; Pillai et al.
2005; Scheibye-Knudsen et al. 2014). Consistent with this, the benefits of NAD+ repletion
through the upregulation of NAD+ biosynthetic enzymes, inhibition of NAD+-consuming
enzymes and pharmacological supplementation with NAD+ precursors can in part be ablated
through deletion of sirtuin activity (Bai et al. 2011; Jang, Kang, and Hwang 2012; Li et al.
2015; Scheibye-Knudsen et al. 2014; Zou et al. 2016). Increasing NAD+ levels may be a
promising strategy to treat some pathologies and age-related diseases (Cantó and Houtkooper
2016; Imai and Guarente 2016; Johnson and Imai 2018; Wątroba and Szukiewicz 2016).
22
Chapter 1 Introduction
The seven mammalian sirtuins each have unique locations and functions—SIRT1, 6
and 7 are localised to the nucleus, SIRT2 is largely a cytoplasmic protein and SIRT3, 4 and 5
are found in the mitochondria. These subcellular localisations are relevant to their activity and
biological function. As describing all the interactions of the sirtuins and their targets is beyond
the scope of this thesis, prominent targets for each of the sirtuins will be discussed below.
SIRT1
SIRT1 is the most well-studied member of the sirtuin family, being closest in homology to the
yeast Sir2 enzyme (Brachmann et al. 1995; Vaziri et al. 2001). SIRT1 is ubiquitously expressed
and is localised largely to the nucleus but can shuttle between the nuclear and cytosolic
compartments in response to cellular stress (Jin et al. 2007b; Michishita et al. 2005; Tanno et
al. 2007). As such, SIRT1 has a range of nuclear and cytosolic targets that favour cell survival
in response to cellular stressors such as nutrient deprivation, oxidative stress, inflammation and
DNA damage.
The first ever identified substrate of SIRT1 was p53 (Luo et al. 2001; Vaziri et al. 2001),
a pro-apoptotic transcription factor and tumour suppressor that, upon deacetylation, inhibits cell cycle arrest and apoptosis (Han et al. 2008; Kim et al. 2007). SIRT1 also plays a role in reducing inflammation by deacetylating nuclear factor kappa B (NF-κB) (Salminen et al.
2008), a family of transcription factors implicated in inflammation, cell death, proliferation and development. This same target is also deacetylated by another nuclear sirtuin, SIRT6, which also dampens inflammation (Kawahara et al. 2009). In response to fasting, SIRT1 acts as a repressor of proliferator-activated receptor gamma (PPARγ) (Picard et al. 2004), a critical regulator of fat storage in white adipose tissue (WAT) and adipocyte differentiation (Rosen and Spiegelman 2006), favouring fat mobilisation instead of storage. A similar effect is demonstrated by cytosolic SIRT2 which deacetylates the transcription factor forkhead box O
1 (FOXO1) (Jing, Gesta, and Kahn 2007), a major regulator of adipocyte differentiation,
23
Chapter 1 Introduction
promoting binding to PPARγ and subsequent repression of its transcriptional activity (Wang
and Tong 2009).
SIRT1 also plays a major role in energy metabolism by deacetylating the transcriptional
coactivator PPARγ coactivator 1-alpha (PGC-1α) (Rodgers et al. 2005), which stimulates
gluconeogenesis, increases fatty acid oxidation and enhances mitochondrial biogenesis
(Dominy Jr et al. 2010; Lagouge et al. 2006). This was also demonstrated in the mitochondrial
compartment where the deacetylation of PGC-1α by SIRT3 enhanced adaptive thermogenesis
and reduced oxidative stress (Giralt et al. 2011; Kong et al. 2010; Shi et al. 2005). SIRT1 is
also important for DNA repair by deacetylating DNA repair proteins such as Ku70 in HR
(Jeong et al. 2007), xeroderma pigmentosum group A (XPA) in NER (Fan and Luo 2010) and
Nijmegen breakage syndrome 1 (NBS1) in aNHEJ (Yuan and Seto 2007). SIRT6 also plays a
role in DNA damage repair by deacetylating DNA polymerase β, the major polymerase
involved in BER (Mostoslavsky et al. 2006). Interestingly, SIRT6, which has both deacetylase
and ADP-ribosyltransferase activities, promotes stress-induced DNA repair (Mao et al. 2011;
Van Meter et al. 2016) and rescues age-related defect in BER (Xu et al. 2015) in a PARP1- dependent manner.
SIRT2
Like SIRT1, the cytosolic SIRT2 can shuttle between the nucleus and cytoplasm in response
to cellular stress such as CR, oxidative stress and inflammation (Michishita et al. 2005).
Upregulation of SIRT2 counteracts stress-induced oxidative stress by binding to and deacetylating the transcription factor FOXO3a. This activates and increases the expression of manganese superoxide dismutase (MnSOD) also known as SOD2 (Kops et al. 2002; Wang et
- al. 2007), a mitochondrial antioxidant which converts superoxide (O2 ) to H2O2 and ultimately
reduces cellular ROS levels (MacMillan-Crow and Thompson 1999; Wang et al. 2007).
24
Chapter 1 Introduction
Deacetylated FOXO3a also promotes its ubiquitination and degradation in a SIRT2-dependent
manner (Wang et al. 2012).
SIRT2 also plays an important role in regulating inflammation by inhibiting the
expression of cytokines (TNF-α, IL-1β, and IL-6), NF-kB and mitogen-activated protein kinase
(MAPK) (Kim et al. 2013b; Rothgiesser et al. 2010). Inhibition of SIRT2 or a lack of SIRT2 activity often leads to detrimental health outcomes. It can cause meiotic arrest, mitochondrial dysfunction and cellular redox imbalance during oocyte maturation (Xu et al. 2019), mediated by an increase in FOXO3a acetylation which blocks nuclear translocation and transcriptional activity (Wang et al. 2017c; Xu et al. 2019). SIRT2 inhibition also exacerbates neuroinflammation and disrupts blood-brain barrier integrity in experimental traumatic brain injury (Yuan et al. 2016). Interestingly, in some cases and through mechanisms that are still unclear, the inhibition of SIRT2 leads to neuroprotection (Chen et al. 2015; Chopra et al. 2012;
Harrison, Smith, and Dexter 2018; Hasegawa et al. 2010; Luthi-Carter et al. 2010; Outeiro et al. 2007; She et al. 2018; Wang et al. 2016a), antidepressant-like action (Erburu et al. 2017;
Muñoz-Cobo et al. 2017), anticancer activity (Hoffmann et al. 2014; Jing et al. 2016; Ma et al.
2014; McCarthy et al. 2013) and anoxia-reoxygenation-induced cell death (Lynn et al. 2008).
SIRT3
The major mitochondrial deacetylase (Lombard et al. 2007), SIRT3 is upregulated in response
to cellular stresses such as CR (Palacios et al. 2009; Qiu et al. 2010; Shi et al. 2005), fasting
(Hirschey et al. 2010), oxidative stress (Qiu et al. 2010) and cold exposure (Shi et al. 2005).
Upregulation of SIRT3 enhances the deacetylation of proteins that protect and regulate the
function and integrity of mitochondria. For example, SIRT3 deacetylates isocitrate
dehydrogenase 2 (IDH2), a mitochondrial enzyme that generates NADPH which is responsible
for regenerating antioxidants GSH (Schlicker et al. 2008; Someya et al. 2010; Yu, Dittenhafer-
Reed, and Denu 2012). Similarly, SIRT3-mediated deacetylation of FOXO3 (Tseng et al. 2014)
25
Chapter 1 Introduction
and MnSOD enhances ROS scavenging activity to reduce oxidative stress in the mitochondria
(Chen et al. 2011b; Qiu et al. 2010; Tao et al. 2010).
SIRT3 is also a major regulator of mitochondrial energy homeostasis by deacetylating
proteins involved in electron transport chain complexes (e.g. NDUFA9 in Complex I) and ATP
synthesis (e.g. ATP synthase F1 proteins) in response to cellular stress (Ahn et al. 2008; Shi et
al. 2005; Vassilopoulos et al. 2014). In a fasted state, SIRT3 stimulates the utilisation of fatty
acids by deacetylating long-chain acyl coenzyme A dehydrogenase (LCAD) and mitochondrial
3-hydroxy-3-methylglutaryl CoA synthase 2 (HMGCS2) generating energy by increasing fatty acid oxidation (Bharathi et al. 2013; Hallows et al. 2011; Hirschey et al. 2010) and ketone body production, respectively (Shimazu et al. 2010). SIRT3 is also important for regulating heat production, stimulating non-shivering thermogenesis in brown adipose tissue by enhancing uncoupling protein (UCP1) expression in the inner mitochondrial membrane. UCP1 is activated by long-chain fatty acids (LCFAs) (Fedorenko, Lishko, and Kirichok 2012) and induces proton leakage-mediated respiration to generate heat instead of ATP (Rossato et al.
2014; Shi et al. 2005; Srivastava et al. 2013).
In the context of cancers, metabolic reprogramming occurs such that there is a shift away from oxidative phosphorylation towards anaerobic glycolysis, known as the Warburg effect (Warburg 1956). SIRT3 represses this by destabilising the transcription factor hypoxia- inducible factor 1 (HIF-1α) (Bell et al. 2011; Finley et al. 2011) which maintains mitochondrial integrity and metabolism under cellular stress (Kim et al. 2010).
SIRT4 SIRT4 is another mitochondrial sirtuin enzyme with ADP-ribosylation and de-lipoylation
(removal of lipoyl group) activity in addition to its deacetylation activity (Ahuja et al. 2007;
Haigis et al. 2006; Laurent et al. 2013; Mathias et al. 2014). One major target of SIRT4 is repressing the activity of enzyme glutamate dehydrogenase (GDH) by ADP-ribosylation which
26
Chapter 1 Introduction prevents insulin secretion in response to amino acids (Ahuja et al. 2007; Haigis et al. 2006).
Interestingly, knock down of SIRT4 increased fatty acid oxidation and mitochondrial function in muscle cells (Nasrin et al. 2010) and consistent with this, another study showed a loss of
SIRT4 increased exercise capacity and protected against diet-induced obesity (Laurent et al.
2013).
In contrast, expression of SIRT4 seems to exhibit a tumour-suppressive function by orchestrating a metabolic block in glutamine metabolism (Jeong et al. 2014; Jeong et al. 2013;
Miyo et al. 2015) with therapeutic potential in the treatment of cancers such as colorectal cancer. There are also discrepancies as to whether SIRT4 activity mediates protection or harm in cardiac function. One study shows SIRT4 protects against hypoxia-induced apoptosis in cardiomyoblast cells (Liu et al. 2013) whereas another study revealed the knockout of SIRT4 offered protection again angiotensin-II induced cardiac hypertrophy and fibrosis in mice (Luo et al. 2017). The role of SIRT4 in response to oxidative stress remains unclear.
SIRT5 In contrast to most other sirtuins, SIRT5 has very weak deacetylase activity (Du et al. 2011). It instead exhibits stronger demalonylation, desuccinylation and deglutarylation activity
(Nakagawa et al. 2009; Nakamura et al. 2012; Park et al. 2013; Polletta et al. 2015). The first of its targets to be identified was carbamoyl phosphate synthetase 1 (CPS1) (Nakagawa et al.
2009), the enzyme catalysing the first step of the urea cycle and plays an important role in detoxification of ammonia. Under nutrient deprived conditions, SIRT5 activity is stimulated and leads to increased desuccinylation of CPS1 and urea formation (Du et al. 2011). SIRT5 has also been shown to improve the antioxidant activity of SOD1 through its desuccinylation, resulting in reduced ROS levels (Lin et al. 2013).
Other targets of SIRT5 are not well understood, though SIRT5 knockout mice exposed to cardiac ischemia exhibited larger infarct volume, increased oxidative stress and fibrosis as
27
Chapter 1 Introduction
well as reduced shortening fraction and ejection fraction (Boylston et al. 2015). An increase in
succinate dehydrogenase (SDH) activity was observed in this study, suggesting that SIRT5 also
desuccinylates SDH and may play a cardioprotective role. Recent studies using metabolomics-
assisted proteomics report SIRT5-deficient mice exhibit defects in fatty acid metabolism,
decreased ATP production and hypertrophic cardiomyopathy, suggesting that SIRT5-mediated
desuccinylation plays a major role in the regulation of heart metabolism and protection
(Sadhukhan et al. 2016).
SIRT6 SIRT6 is a nuclear sirtuin with deacetylation, ADP-ribosylation, and de-myristoylation activity. SIRT6 is required for the maintenance of glucose and lipid homeostasis by co- repressing the transcription factor hypoxia-inducible factor 1-alpha (HIF-1α) (Zhong et al.
2010) which in turn supresses glucose uptake and glycolysis. SIRT6 also plays a major role in genomic stability and DNA integrity. One of its targets is the histone H3 lysine 9 (H3K9) which modulates telomeric chromatin (Michishita et al. 2008). Further, activation of SIRT6 promotes
DNA repair under oxidative stress by binding to and ADP-ribosylating PARP1, hence
stimulating its DNA repair capacity (Mao et al. 2011). SIRT6 deacetylates C-terminal binding
protein (CtBP) interacting protein (CtIP), which promotes resection on either side of the DNA
break which is critical for DNA repair via HR (Kaidi et al. 2010). Interestingly, the
overexpression of Sirt6 demonstrated significantly longer lifespans in male, but not female
transgenic mice with lower serum IGF1 levels and altered IGF1 signalling, a key pathway in
the regulation of lifespan (Kanfi et al. 2012).
SIRT7 Finally, the last nuclear sirtuin, SIRT7 is found in the nucleolus and interacts with RNA
polymerase I (RNA Pol I), increasing RNA Pol I-mediated transcription involving cell proliferation and apoptosis. SIRT7-deficient mice experience a reduced median and maximum
28
Chapter 1 Introduction
lifespan and develop heart complications including cardiac hypertrophy, inflammatory
cardiomyopathy and extensive fibrosis that was related to hyperacetylation of p53
(Vakhrusheva et al. 2008). The same study showed that SIRT7 deficient cells also exhibit a
~200% increase in apoptosis and severely diminished resistance to oxidative stress, indicating
that SIRT7 plays a major role in the regulation of stress response and cell death in the heart.
Another target of SIRT7 is the transcription factor Myc which plays a role reducing
endoplasmic reticulum (ER) stress upon SIRT7-mediated deacetylation and can prevent fatty
liver disease (Shin et al. 2013).
Regulation of sirtuin activity
A prominent hypothesis in the field is that maintaining sirtuin activity may be needed to
maintain health and protection against conditions such as type 2 diabetes and metabolic
syndrome (Banks et al. 2008; Caron et al. 2014; Halperin-Sheinfeld et al. 2012; Herranz et al.
2010). Understanding the mechanisms that regulate their levels of expression and transcriptional activity may be relevant to the study of age-related diseases. Beyond their
activation in response to cellular stress or during conditions that compromise nutrient and
energy availability in the cell such as CR (Cohen et al. 2004; Nemoto, Fergusson, and Finkel
2004; Noriega et al. 2011), the sirtuins are also regulated at the level of transcription, post-
translational modification, through protein-protein interactions or by levels of its substrate
NAD+.
At the level of transcription, CR increases the protein expression and activity of SIRT1
(Cohen et al. 2004; Civitarese et al. 2007), SIRT3 (Lombard et al. 2007; Schwer et al. 2009)
and SIRT5 (Nakagawa et al. 2009). Exercise also increases SIRT1 mRNA levels in skeletal
muscle and SIRT3 protein levels increase in the triceps of inactive young men (Radak et al.
2011). Multiple transcriptional binding sites have been identified where several transcription
factors bind to and regulate SIRT1 expression and transcription in response to low energy status
29
Chapter 1 Introduction such as CR and exercise. For example, under normal energy status, the pro-apoptotic transcription factor and tumour suppressor p53 is bound to the SIRT1 promoter, repressing
SIRT1 gene expression. During situations of bio-energetic stress, p53 bound to the SIRT1 promoter is removed, a process that relies on the translocation of FOXO3a from the cytoplasm
(inactive) to the nucleus (active) to form a complex with p53 (Nemoto, Fergusson, and Finkel
2004). Interestingly, p53 is deacetylated directly by SIRT1, destabilising and inactivating its tumour suppressing functions and also promoting the transcription of SIRT1 that is normally repressed under basal conditions (Chen et al. 2005; Yamakuchi and Lowenstein 2009).
The expression of SIRT1 can also be regulated and repressed by another tumour suppressor, hypermethylated in cancer 1 (HIC1) in response to DNA damage (Chen et al.
2005). HIC1 deletion results in an increase in SIRT1 transcription and subsequent deacetylation and inactivation of p53, maintaining cell survival during DNA damage (Chen et al. 2005).
HIC1 can also repress SIRT1 by forming a complex with CtBP, which is activated by elevated
NADH levels and in turn leads to higher binding affinity with HIC1. Changes to cellular redox status, such as when glycolysis is blocked, alters the binding of CtBP to HIC1 which in turn reduces HIC1 binding and repression of SIRT1, inducing SIRT1 expression and transcription during CR (Zhang et al. 2007).
The DNA damage repair enzyme PARP2 is also a transcriptional repressor of SIRT1, where SIRT1 mRNA expression is increased in PARP2-deficient myotubes, suggesting
PARP2 is a direct negative regulator of the SIRT1 promotor (Bai et al. 2011). Consistent with the increase in sirtuin expression and activity under low cellular energy status, exposure to conditions that overload the cell with nutrients and energy, such as through the consumption of a high-fat diet, reduces mRNA and protein levels of SIRT1 in adipose tissue of mice
(Chalkiadaki and Guarente 2012) and these levels are also implicated in the subcutaneous adipose tissue of obese individuals (Pedersen et al. 2008).
30
Chapter 1 Introduction
A potential reason for low SIRT1 expression and activity could be the reduced
bioavailability of its substrate NAD+, which declines in livers and white adipose tissues of
high-fat-diet-fed mice (Yoshino et al. 2011). This same study showed that supplementation
with NAD+ precursor NMN restored NAD+ levels and ameliorated the pathophysiology associated with the consumption of a high-fat-diet in a SIRT1-dependent manner (Yoshino et
al. 2011). These studies highlight the importance of the coordination between energy-sensing
enzymes and transcription factors in regulating sirtuin expression, transcription and activation
in response to cellular stress.
One of the ways in which the sirtuins can be regulated at the post-translational level is through phosphorylation (Sasaki et al. 2008). SIRT1 has 15 phosphorylation sites, which affects their activity and regulates their own protein levels by influencing protein degradation pathways (Sasaki et al. 2008; Zschoernig and Mahlknecht 2009). SIRT1 can be phosphorylated by protein kinases including c-Jun N-terminal kinases (JNKs), casein kinase 2 (CK2), Cyclin
B/cyclin-dependent kinase 1 (Cyclin-B/Cdk1), and the dual specificity tyrosine phosphorylation-regulated kinases (DYRKs). Together, these phosphorylation events increase
SIRT1 deacetylation activity to counteract oxidative stress (Nasrin et al. 2009), reduce DNA damage (Kang et al. 2009), allow cell cycle progression and proliferation (Sasaki et al. 2008) and promote cell survival by inhibiting apoptosis (Guo et al. 2010).
SIRT2 can also be phosphorylated by kinases such as Cyclin E/Cdk2 which inhibits its deacetylase activity, blocking cell cycle progression and arresting cells in the G1 phase
(Pandithage et al. 2008). Another post-translational modification to the sirtuins is through the addition of small ubiquitin-like modifier (SUMO), otherwise known as sumoylation (Yang et al. 2007b). Under basal conditions, sumoylated SIRT1 can deacetylate and inactivate p53 whereas in response to oxidative stress and DNA damage, SIRT1 is desumoylated, and p53 is activated to induce cell death (Yang et al. 2007b).
31
Chapter 1 Introduction
The protein deleted in breast cancer 1 (DBC1) directly binds to and suppresses SIRT1
activity. When treated with the chemotherapeutic drug etoposide, DBC1 associates with
SIRT1, inhibiting its activity and leading to increased acetylation of p53 and its pro-apoptotic
functions (Kim, Chen, and Lou 2008; Zhao et al. 2008). The expression of DBC1 is associated
with a poorer prognosis in certain cancers (Cha et al. 2009; Kim et al. 2013a; Lee et al. 2011;
Noh et al. 2013), suggesting its inhibition, rather than expression, as a new therapeutic target
in the treatment of cancer. A recent study showed that the metastasis suppressor breast cancer
metastasis suppressor 1 (BRMS1) interferes with DBC1-SIRT1 interaction by binding to
DBC1 and inducing SIRT1-mediated p53 deacetylation under genotoxic stress (Liu et al.
2016). More studies are needed to understand the role of BRMS1 in DBC1-mediated inhibition
of SIRT1 deacetylation activity and how it can be used in the treatment of cancers.
As the activity of the sirtuins is dependent on the binding of NAD+ to their catalytic site, NAD+ levels are another factor regulating sirtuin activity. NAD+ is used as a cofactor in
redox reactions and are also consumed by other enzymes such as PARPs and CD38/CD157,
and increased NAD+-consuming activities can cause a rapid decline in NAD+ levels (Camacho-
Pereira et al. 2016; Wang et al. 2013), potentially compromising the activity of the sirtuins and
other NAD+ dependent enzymes. Raising NAD+ levels by increasing the expression of NAD+
biosynthetic enzymes (Yang et al. 2007a), pharmacologically inhibiting NAD+-consuming enzymes (Bai et al. 2011; Cerutti et al. 2014; Tarragó et al. 2018) or pharmacologically supplementing with NAD+ (Wang et al. 2013) or NAD+ precursors (Cerutti et al. 2014; Das et
al. 2018; Wang et al. 2018; Zou et al. 2016) leads to increased activity of the sirtuins, and is
associated with an improvement in health and disease parameters.
The sirtuins are also regulated by NAM, the by-product of NAD+ hydrolysis, creating
a negative feedback loop (Bitterman et al. 2002), though this inhibition only occurs at very
high doses, in the millimolar range. Binding of NAM is non-competitive and occurs at a
32
Chapter 1 Introduction
conserved pocket adjacent to NAD+, which blocks the binding of NAD+ and subsequent hydrolysis. Supplementation with exogenous NAM can inhibit SIRT1-mediated deacetylation of p53, which inhibits apoptosis and in the context of cancers has been shown to block proliferation and promote apoptosis in cancerous cells (Audrito et al. 2011). The application of exogenous NAM has also been shown to mediate anti-viral effects by suppressing SIRT1- mediated enhancement of viral DNA replication (Li et al. 2016).
1.2.2.3 NAD+ and CD38/CD157
Cluster of differentiation 38 (CD38) is a cell surface enzyme highly expressed in immune cells
such as lymphocytes, monocytes, macrophages, dendritic cells, granulocytes and natural killer
(NK) cells (Quarona et al. 2013). Together with its homologue CD157 (also known as BST1;
bone marrow stromal cell antigen 1), CD38 has several different enzymatic activities in various
biological roles.
CD38 and CD157 act as ADP-ribose cyclases which hydrolyse NAD+ to form NAM
and ADPR and cyclic ADPR (cADPR). NAM can be recycled as a substrate for synthesis back
into NAD+. ADPR is a source of extracellular adenosine, an important molecule generated in
response to cellular stress and plays an essential role in almost all aspects of physiology (Borea
et al. 2018). cADPR plays an important role as a secondary messenger that mobilises the release
of intracellular calcium (Ca2+) to regulate physiological processes including the immune
response, nervous system excitability, muscle contraction, enzyme activity and circadian
rhythm (Dodd et al. 2007; Wang et al. 2004; White, Kannan, and Walseth 2003).
CD38 is also responsible for the generation of another secondary messenger known as
nicotinic acid adenine dinucleotide phosphate (NAADP), which is produced through a base-
exchange reaction between NADP+ and nicotinic acid (NA) in an acidic environment (Aarhus
et al. 1995; Chini and Dousa 1995; Liang et al. 1999). NAADP regulates calcium/sodium
33
Chapter 1 Introduction
(Ca2+/Na+) channels (known as two pore calcium channels; TPCN1 and 2) (Morgan et al. 2015;
Ruas et al. 2015) which promotes calcium release from lysosomes (Fang et al. 2018) and inhibits autophagy (Lu et al. 2013b). Further, TPCN channels are important regulators of the mTOR pathway (Cang et al. 2013). As both autophagy and mTOR are dysregulated in ageing, it is possible that NAADP, through its regulation of TPCN channels, plays an important role in the ageing process through autophagy-/mTOR-mediated pathways.
For many years, it was thought that the main function of CD38 was to generate the secondary messengers cADPR and NAADP, though this remains controversial for several
reasons. First, the generation of cADPR is a highly inefficiency process, requiring nearly 100
molecules of NAD+ to generate 1 molecule of cADPR (Zielinska et al 2004). Second, though
NAADP synthesis has been shown to occur in cells overexpressing CD38 (Cosker et al. 2010),
it is unknown whether NAADP is generated in cells naturally expressing CD38 or in vivo.
Further, NAADP can be generated in the absence of CD38 expression (Soares et al. 2007),
indicating there are CD38-independent pathways to synthesise NAADP (Kim et al. 2008).
CD38 is a major consumer and key regulator of intracellular NAD+ levels (Aksoy et al.
2006; Barbosa et al. 2007). Pharmacokinetic studies have shown the NAD+-hydrolysing
efficiency of CD38 is several-fold higher than its homologue CD157 (Hussain, Lee, and Chang
1998) and compared to other NAD+-consuming enzymes, CD38 has one of the lowest binding
+ affinities for NAD with a Michaelis-Menten constant (Km) of 15-25 µM versus SIRT1-7
(range ~13-888 µM), PARP1 (20-97 µM), PARP2 (130 µM) and SARM1 (24 µM) (Katsyuba et al. 2020), though other work has suggested that these Km values are well below physiological
NAD+ levels of 400-700 µM, and may not be sensitive to physiological changes in NAD+ (Hara et al. 2019). Inhibition of CD38 could therefore increase overall NAD+ levels (Aksoy et al.
2006; Tarragó et al. 2018).
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Chapter 1 Introduction
Interestingly, while the majority of CD38 acts as an ecto-NADase, hydrolysing NAD+
in the extracellular compartment, NAD+ is mostly intracellular. This topological paradox is
explained by recent discoveries that CD38 can be expressed in two opposing membrane
orientations with one catalytic site facing the outside (type II) and another facing the cytoplasm
(type III) (Liu et al. 2017a; Zhao, Lam, and Lee 2012). And, in addition to NAD+, CD38 also hydrolyses the extracellular NAD+ precursors NMN and nicotinamide riboside (NR) prior to
their transport into the cell (Grozio et al. 2013; Camacho-Pereira et al. 2016; Preugschat et al.
2014). These findings add weight to the notion that CD38 plays a critical role in the
biosynthesis and metabolism of intracellular NAD+ levels by hydrolysing its extracellular
precursors. Further, pools of intracellular CD38 have been found embedded in the inner
membranes of the nucleus and mitochondria and likely as a soluble form in the cytoplasm
(Chini 2009; Hogan, Chini, and Chini 2019; Malavasi et al. 2008; Shrimp et al. 2014),
demonstrating multiple routes for CD38 to access and degrade intracellular NAD+.
CD38 also plays a role in the generation of extracellular adenosine, a nucleoside
involved in regulating processes such as inflammation and the immune response, neuronal
activity, vascular function, platelet aggregation and blood cell regulation (Borea et al. 2018).
CD38 forms an axis with the ecto-nucleotide pyrophosphatase/phosphodiesterase 1 (NPP1,
also known as CD203a or PC-1) and the ecto-5’-nucleotidase CD73, independent of the
nucleoside tri- and diphosphohydrolase (NTPDase) CD39, which generates extracellular adenosine from ATP (Horenstein et al. 2013). CD38 hydrolyses extracellular NAD+, NMN or
NR forming ADPR which in turn is broken down into adenosine monophosphate (AMP) via
NPP1, followed by CD73 which catalyses the conversion of AMP to adenosine. Adenosine
mediates its effects by activating G protein-coupled receptors expressed on the cell membrane
(Borea et al. 2018). It is generated in response to cellular stresses such as hypoxia, metabolic stress or injury and promotes downstream processes required to prevent further cellular
35
Chapter 1 Introduction
damage. The upregulation of CD38-mediated adenosine receptor signalling causes
immunosuppression during cancer therapy by inhibiting CD8+ T-cell function, and was reported as a mechanism for tumour cells to escape detection through immune checkpoints such as programmed death 1 (PD-1) and its ligand, PD-L1 (Horenstein et al. 2013; Nijhof et
al. 2016). This has also been implicated in other cancers such as multiple myeloma (Horenstein
et al. 2016) and CD38 inhibition has emerged as an effective target to overcome resistance to
immune checkpoint inhibitors in cancer immunotherapy (Chen et al. 2018).
CD38 activity is also upregulated in normal biological ageing, which is has been
correlated with declining NAD+ levels (Camacho-Pereira et al. 2016; Schultz and Sinclair
2016). Preventing CD38-induced NAD+ depletion through the generation of CD38-deficient
mice or supplementing with CD38 inhibitors increased NAD+ levels up to 30-fold (Aksoy et
al. 2006; Barbosa et al. 2007; Young et al. 2006) and similar studies show it protects against
pathological conditions such as age-related metabolic dysfunction (Tarragó et al. 2018), postischemic heart dysfunction (Boslett et al. 2017) and the progression of cancers such as glioma (Blacher et al. 2015).
The cellular distribution of CD38, taken together with its various topological forms and cellular location suggests that this enzyme may play a role in NAD homeostasis. While upregulating CD38 activity depletes the cell of the crucial metabolite NAD+, it is also a key generator of important secondary messengers such as cADPR, ADPR, NAADP and adenosine.
Under certain conditions, inhibiting CD38 activity may have detrimental outcomes (Xu et al.
2012; Zhang et al. 2014b). Understanding the conditions under which the activation or inhibition of CD38 activity is beneficial, versus detrimental, is therefore important when considering how it may affect its downstream pathways including adenosine signalling, calcium mobilisation and NAD+ metabolism.
36
Chapter 1 Introduction
2.2.4 NAD+ and SARM1
Finally, the central executioner of the axonal degeneration pathway, sterile alpha and
Toll/interleukin-1 receptor motif-containing 1 (SARM1) (Gerdts et al. 2013; Osterloh et al.
2012) was recently shown to possess intrinsic NAD+ cleavage activity causing injury-induced
loss of NAD+ in neurons (Essuman et al. 2017; Gerdts et al. 2015; Summers et al. 2016).
SARM1-induced depletion of NAD+ triggers axon degeneration which can be blocked by
overexpressing NAD+ biosynthetic enzymes such as NAMPT and NMNAT (Gerdts et al. 2015;
Sasaki et al. 2016), or through pharmacological supplementation with NAD+ precursors such
as NR (Gerdts et al. 2015). Preventing the loss of NAD+ through SARM1 deletion (Gilley et
al. 2015; Gilley, Ribchester, and Coleman 2017; Turkiew et al. 2017) or supplementation with
the SARM1 inhibitor XAV939 (Gerdts et al. 2015) rescued these neuronal defects and axon
degeneration.
Interestingly, accumulation of NMN, the direct precursor for NAD+ biosynthesis via
NMNAT, seems to promote axon degeneration and the neuroprotective effects of NMNAT
enzymes may be due to its consumption of NMN, rather than its synthesis of NAD+ (Di Stefano
et al. 2015). This was first noted in the Wallerian degeneration Slow (WldS) mutant mouse
model, which encodes an intact and functional NMNAT1 enzyme and resists axon
degeneration but showed no difference in NAD+ levels in a WldS mouse brain (Mack et al.
2001). Recent studies have supported the notion that the protective effect of WldS mice and
NMNAT overexpression is not due to increasing NAD+ levels by showing transient expression of bacterial NMN deamidase in mice, which has NMN-consuming without NAD+-synthesising
activity, delays axon degeneration (Di Stefano et al. 2015; Di Stefano et al. 2017). Further, the
degenerative effects of NMN accumulation was recently shown to stimulate SARM1 activity
by ~3.5-fold, causing NAD+ depletion-induced non-apoptotic cell death (Zhao et al. 2019), and
more studies on this are warranted.
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Chapter 1 Introduction
As the first member of a new class of NAD+-consuming enzymes, loss of SARM1
activity has emerged as an attractive therapeutic target to treat various neuropathies including
axon degeneration (Geisler et al. 2019) and traumatic brain injury (Henninger et al. 2016;
Marion, McDaniel, and Armstrong 2019), as well as neurodegenerative diseases such as
Alzheimer’s, Parkinson’s and Huntington’s disease (Krauss et al. 2020).
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Chapter 1 Introduction
1.3 NAD+ deficiency is a critical pathological factor in ageing and disease
1.3.1 NAD+ deficiency in ageing and disease
There is mounting evidence from rodent and human studies showing NAD+ levels decline with
age in skeletal muscle (Mendelsohn and Larrick 2014; Yoshino et al. 2011), brain (Braidy et
al. 2014; Stein and Imai 2014; Zhu et al. 2015), heart (Braidy et al. 2011a; Pillai et al. 2005),
lung (Braidy et al. 2011a), liver (Braidy et al. 2011a; Zhou et al. 2016), kidney (Braidy et al.
2011a), pancreas (Yoshino et al. 2011), white adipose tissue (Yoshino et al. 2011), skin
(Massudi et al. 2012a), endothelial cells (Das et al. 2018) and oocytes (Bertoldo et al. 2020).
NAD+ deficiency is a common pathological factor implicated in a number of disease models. These include cardiometabolic diseases such as obesity (Choi et al. 2013), type 2 diabetes (Yoshino et al. 2011), non-alcoholic fatty liver disease (Gariani et al. 2016), myocardial I/R injury (Yamamoto et al. 2014a) and cardiac hypertrophy (Pillai et al. 2005).
Preclinical efficacy has also been shown in neurological disorders such as cerebral
ischemia/reperfusion (I/R) injury (Liu et al. 2009; Park et al. 2016), traumatic brain injury
(Clark et al. 2007; Satchell et al. 2003), Ataxia Telangiectasia (AT) (Fang et al. 2016),
Parkinson’s disease (PD) (Lu et al. 2014a), and Alzheimer’s disease (AD) (Gong et al. 2013).
Finally, this strategy has also been implicated in skeletal muscle degeneration in a mouse model of Duchene’s muscular dystrophy (Ryu et al. 2016) and chemotherapy- and inflammation- induced tissue injury (Tateishi et al. 2017; Umapathy et al. 2012). Collectively, these studies show that NAD+ deficiency is a critical pathological factor in ageing and disease and is a
promising strategy to potentially treat age-related diseases and the overall physiological decline with age.
39
Chapter 1 Introduction
1.3.2 Increased NAD+ consumption as a cause of NAD+ deficiency
Declining NAD+ levels in ageing and disease is attributed to increased NAD+ consumption and/or decreased NAD+ biosynthesis. The major consumers of NAD+ in the cell are the PARPs
(Alano et al. 2010) and CD38 (Camacho-Pereira et al. 2016), whose activities also increase in
an age-dependent manner and correlate with declining NAD+ levels with age (Camacho-Pereira et al. 2016; Massudi et al. 2012b). Activation of PARP1 in rodent and human models of ageing is predominantly triggered by increased DNA damage (Braidy et al. 2011b; Braidy et al. 2014;
Fang et al. 2014; Massudi et al. 2012b) which, when extensive or ineffectively repaired, lead to the hyperactivation of PARP1 followed by the rapid depletion of NAD+ (Fang et al. 2014;
Zhang et al. 2014a; Zhou et al. 2006).
NAD+ deficiency impedes proper functions of the sirtuins and PARPs, leading to
mitochondrial dysfunction (Fang et al. 2014; Scheibye-Knudsen et al. 2014; Zhou et al. 2006),
a major hallmark of ageing (López-Otín et al. 2013), and cell death (Alano et al. 2010; Muñoz-
Gámez et al. 2009; Zhang et al. 2014a). This is demonstrated in a recent study using a mouse
model of the accelerated ageing disorder Cockayne syndrome (CS) which is characterised by
progressive neurodegeneration due to mutations in genes encoding the DNA repair proteins CS
group A (CSA) or CS group B (CSB) (Scheibye-Knudsen et al. 2014). CSB deficiency in these
mice led to PARP1 activation, NAD+ depletion and attenuation of SIRT1 activity causing
mitochondrial dysfunction, inflammation and cell death. In this model, treatment with PARP1
inhibitors could rescue CS-associated phenotypes by restoring NAD+ and activating SIRT1
(Scheibye-Knudsen et al. 2014).
NAD+ deficiency can also be caused by increased CD38 activity with age, which attenuates SIRT3 activity and leads to mitochondrial dysfunction, characterised by abnormal mitochondrial morphology, reduced mitochondrial oxygen consumption rate and a significant reduction in ATP synthesis (Camacho-Pereira et al. 2016). CD38 gene expression may be
40
Chapter 1 Introduction
upregulated by inflammatory cytokines secreted by senescent cells as part of the senescent
associated secretory phenotype (SASP) (Chini et al. 2019), another major hallmark of ageing
(López-Otín et al. 2013) or bacterial endotoxins such as lipopolysaccharide (LPS) (Lee et al.
2012; Shu et al. 2018), contributing to cellular senescence and inflammation frequently
observed during ageing and age-related diseases (Franceschi and Campisi 2014; He and
Sharpless 2017).
Treatment with CD38 inhibitors (PMID 25828863)or genetic deletion of CD38 can
raise NAD levels, and have been effective in the treatment of pathological conditions such as
acute kidney injury (Shu et al. 2018), ischemic brain damage (Long et al. 2017)and cardiac
hypertrophy (Guan et al. 2017a). A note of caution, however, is warranted as CD38 catalyses
the synthesis of cADPR and its inhibition or genetic deletion may have deleterious impacts on
other cADPR-mediated roles. For example, cADPR is involved in the regulation of the immune
response (Frasca et al. 2006) as well as the release of oxytocin which controls many social
behaviours (Jin et al. 2007a). Inhibiting CD38 may increase susceptibility to bacterial
infections, weaken the immune response and induce mental impairments in individuals.
In contrast to the increase in activity of PARPs and CD38 with age, the activity of the
sirtuins decline with age (Braidy et al. 2011b; Massudi et al. 2012b), which has been attributed
to PARP- and CD38-mediated NAD+ depletion (Camacho-Pereira et al. 2016; Pillai et al. 2005;
Wang et al. 2013). Controversy surrounds this idea, as the affinity of sirtuins for NAD+ is below
the concentrations seen in the age-related decline in NAD+ availability, and it is not clear
whether declining NAD+ is related to declining sirtuin activity. Despite their extensive roles to
maintain cellular homeostasis, the activity of the sirtuins contribute very little to the depletion
of intracellular NAD+ stores, likely due to their tight regulation in the cell by several different mechanisms (Section 2.2.2 NAD+ and the sirtuins) including the generation of NAM from
NAD+ hydrolysis, which is an inhibitor of the sirtuins.
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Chapter 1 Introduction
1.3.3 Decreased NAD+ biosynthesis as a cause of NAD+ deficiency
In addition to increased NAD+-consumption, NAD+ biosynthetic activity declines with age.
The rate-limiting enzyme for NAD+ biosynthesis in the salvage pathway, NAMPT, catalyses
the conversion of NAM to NMN (Revollo, Grimm, and Imai 2004) and is an important
regulator of the intracellular NAD+ pool (Garten et al. 2015). The decline in NAD+ with ageing has been heavily associated with the downregulation of this enzyme in several bodies of work.
For example, NAMPT levels decline with age in the hippocampus, where NAMPT is the main source of NAD+ biosynthesis for adult neural/progenitor stem cells (NPSC), crucial for proper cognitive function (Stein and Imai 2014). NSPC-specific loss of NAMPT was associated with a reduced capacity to proliferate, self-renew and differentiate into oligodendrocytes, leading to a decline in cognitive function (Stein and Imai 2014). Similarly, NAMPT expression was
decreased in mesenchymal stem cells obtained from aged rats (Ma et al. 2017) and in retinal
pigment epithelium from aged mice (Jadeja et al. 2018) compared to young controls. Reduced
NAD+ levels in these cells was associated with decreased SIRT1 activity and enhanced cellular
senescence, a major hallmark of ageing (López-Otín et al. 2013).
The consequences of NAD+ deficiency due to a loss of NAMPT expression were also
demonstrated in mouse models where the genetic deletion of Nampt in adult projection neurons
led to motor dysfunction and death (Wang et al. 2017b). Similarly, adipocyte-specific loss of
Nampt led to the development of multi-organ insulin resistance and impairment in adipose tissue functions (Stromsdorfer et al. 2016) whereas in skeletal muscle, Nampt deletion caused
muscle fibre degeneration, loss of muscle strength and reduced treadmill endurance (Frederick
et al. 2016). These studies demonstrate the importance of NAMPT in preserving the proper
functional capacity of tissues and preventing systemic metabolic complications by maintaining
adequate levels of NAD+ in cells. Supplementation with NAD+ precursors such as NMN and
NR, which synthesise NAD+ via NMNAT (Berger et al. 2005) and NR kinases (NRK1 or
42
Chapter 1 Introduction
NRK2), bypass the rate-limiting step (Tempel et al. 2007) and could ameliorate the functional
and metabolic defects in these studies by restoring NAD+ levels.
Apart from NAMPT, overexpression of enzymes responsible for NAD+ biosynthesis in the mitochondria, NMNAT3 and NNT (Section 2.1.2 NADP+ and NADPH roles and functions), restored age-dependent decline in mitochondrial NAD+ levels and increased the
activity of the mitochondrial sirtuin, SIRT3. This restored redox homeostasis, protected against
oxidative stress-induced cellular damage, enhanced reprogramming efficiency and led to the
extension of lifespan of mesenchymal stem cells by delaying replicative senescence (Son et al.
2016). Overall, these studies suggest that NAD+ deficiency, either due to increased
consumption or decreased biosynthesis, may be a critical factor in ageing and disease, and
strategies to restore NAD+ levels represent a potential therapeutic target to treat age-related
physiological decline and age-related diseases.
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Chapter 1 Introduction
1.4 Consequences of NAD+ deficiency in cell function and survival
Due to the significant involvement of NAD+ in hundreds of reactions in the cell, declining
NAD+ levels can cause extensive impairments in cellular function and threaten cell survival.
A major consequence of NAD+ deficiency is mitochondrial dysfunction, a major
hallmark of aging (López-Otín et al. 2013). Mitochondrial dysfunction caused by low NAD+
levels, is commonly characterised by increased mitochondrial permeability transition (MPT),
release of proapoptotic factors such as apoptosis-inducing factor (AIF) and blockage of
glycolysis, leading to reduced ATP production (Camacho-Pereira et al. 2016; Di Lisa et al.
2001; Ha and Snyder 1999; Ying, Garnier, and Swanson 2003; Yu et al. 2002). This can cause the cell to enter an energy crisis and ultimately lead to cell death (Alano et al. 2010; Di Lisa et al. 2001; Wang et al. 2013; Ying, Garnier, and Swanson 2003).
Mitochondrial dysfunction may also induce cellular senescence, another major hallmark of ageing (López-Otín et al. 2013), and mitochondrial dysfunction-associated senescence reduced NAD+/NADH ratios and accelerated the ageing process (Wiley et al.
2016). The link between declining NAD+ levels and mitochondrial dysfunction has in part been
attributed to reduced activity of the sirtuins, which play a major role in the maintenance of
mitochondrial function. For example, the decline in NAD+ levels with age is associated with
reduced SIRT1 activity, impeding its ability to control the expression of mitochondria-encoded proteins, which are required for the coordination of and communication between the nuclear and mitochondrial genomes (Gomes et al. 2013).
Similarly, increased CD38 activity with age is correlated with a severe decline in NAD+
levels, reduced SIRT3 activity and mitochondrial dysfunction, whereas genetic deletion of
CD38 in mice led to an increase in NAD+ levels, increased SIRT3 activity and improved
mitochondrial function (Camacho-Pereira et al. 2016). These findings suggest that
44
Chapter 1 Introduction
mitochondrial function is highly dependent on SIRT1 and SIRT3 activity but require adequate
levels of intracellular NAD+ for proper function and protection. Restoring NAD+ levels by
inhibiting NAD+-consuming enzymes such as PARPs or supplementing with NAD+ precursors such as NMN and NR significantly attenuated oxidative stress-induced mitochondrial dysfunction, AIF release and neuronal cell death (Du et al. 2003; Sims et al. 2018), improving the pathophysiology of animal models of disease caused by mitochondrial dysfunction. This includes a mouse model of Ataxia Telangiectasia (Fang et al. 2016) and heart failure.
Another major consequence of NAD+ deficiency is reduced DNA repair capacity due
to reduced activity of the DNA repair enzymes such as PARPs and sirtuins (Braidy et al. 2011b;
Massudi et al. 2012a) which have been implicated in mouse models of disease such as Ataxia
Telangiectasia (Fang et al. 2016), xeroderma pigmentosum group A (XPA) (Fang et al. 2014),
Cockayne syndrome (Scheibye-Knudsen et al. 2014) and Alzheimer’s disease (Hou et al.
2018). Inhibition of PARPs and supplementation with NAD+ precursors enhanced the DNA damage repair capacity in these studies by stimulating SIRT1 and SIRT3, highlighting a role in NAD+ augmentation therapies to treat accelerated ageing syndromes and neurodegeneration
by enhancing activity of the sirtuins.
Finally, NAD+ deficiency may contribute to global cellular damage in the form of
increased oxidative stress (Braidy et al. 2011b; Massudi et al. 2012b), inflammation (Minhas
et al. 2019) and cell death (Zhou et al. 2015; Zhu et al. 2016). The accumulation of these process
has been associated with the development and progression of diabetes (Ogura et al. 2018;
Yoshino et al. 2011), acute kidney injury (Mehr et al. 2018) and neurodegeneration (Zhou et
al. 2015). NAD+ augmentation strategies, through administration of CD38 inhibitors or NAD+
precursors, significantly improved the pathophysiologies associated with diabetes, neuronal
and renal dysfunctions and highlight their therapeutic potential to treat age-related diseases by
reducing oxidative stress, inflammation and cell death.
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Chapter 1 Introduction
1.5 Therapeutic potential of NAD+ boosting strategies
NAD+ is synthesised from precursors including NA, NAM, NR and NMN. These can be
sourced naturally from the dietary intake of vegetables, fruits, meat and milk (Krehl et al. 1946;
Mills et al. 2016; Trammell et al. 2016d; Ummarino et al. 2017) which is sufficient to prevent the development of pellagra, a disease caused by NAD+ deficiency and characterised by
dermatitis, diarrhea, and dementia (Hegyi, Schwartz, and Hegyi 2004). While dietary intake of
these precursors can prevent chronic NAD+ insufficiency, it is not clear that dietary sources
alone can correct the declining levels of NAD+ observed during biological ageing and disease.
Exogenous treatment with NAD+ precursors is an attractive strategy for treating age-related
disease, and treatment with NA is an FDA approved therapeutic for cardiovascular disease in
humans (Brown and Zhao 2008).
Studies in humans and rats have shown that NA is the most effective agent available in
the treatment of hypercholesterolemia (Davignon et al. 1994; Shah et al. 2013), though to avoid
painful flushing induced by its interaction with the G-protein coupled receptor GPR109a, is delivered clinically as a slow-release tablet (Benyó et al. 2005). While several rodent studies show NAM provides protection against conditions such as diabetes, glaucoma and renal dysfunction (John et al. 2012; Mitchell et al. 2018; Williams et al. 2017; Zheng et al. 2019), it is a sirtuin inhibitor (Bitterman et al. 2002). Supplementation with NAM at high doses can lead to undesirable or non-beneficial health outcomes, potentially through blocking sirtuin activity
(Li et al. 2013; Smith et al. 2019). In contrast to NA and NAM, the NAD+ precursors NMN
and NR can effectively raise NAD+ in cells without causing painful flushing or inhibiting the
sirtuins, and can be tolerated at high doses and administered for long periods without presenting
harmful side effects in mice for NMN (Mills et al. 2016) and in humans for NR (Martens et al.
2018).
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Chapter 1 Introduction
There are, however, exceptions to these studies though they demonstrate largely
beneficial effects from NMN and NR treatment. One study using cultured superior cervical
ganglia (SCG) explants, neurites and cell bodies from mice showed that exogenous NMN can
promote axon degeneration, which can be rescued by inhibiting its synthesis via FK866
treatment or through over-expression of the NMN-consuming enzyme NMNAT3 (Di Stefano
et al. 2015). Another study showed that NR treatment reduced exercise performance in rats
(Kourtzidis et al. 2016), however this contrasts with findings from another study which show
that NMN treatment enhances physical endurance in aged mice (Das et al. 2018). The
mechanisms through which these contradictory findings occur are not completely understood
and a majority of preclinical rodent studies using NMN and NR have consistently reported
significant improvements in healthspan and lifespan (Rajman, Chwalek, and Sinclair 2018;
Yoshino, Baur, and Imai 2018).
NMN administration in preclinical rodent models of disease have shown to improve
pathological conditions including Alzheimer’s disease in mice (Yao et al. 2017), cerebral
ischemia in mice (Park et al. 2016), cognitive impairments in mice (Tarantini et al. 2019),
obesity in mice (Uddin et al. 2017), diabetes in mice (Yoshino et al. 2011), acute kidney injury
in mice (Guan et al. 2017b), myocardial I/R injury in mice (Yamamoto et al. 2014b), heart
failure in mice (Zhang et al. 2017), haemorrhagic shock in rats (Sims et al. 2018), age-related physiological decline in mice (Mills et al. 2016) and age-related infertility in mice (Bertoldo et al. 2020). Similarly, NR has demonstrated therapeutic potential in the treatment of preclinical disease models including obesity in mice (Cantó et al. 2012), diabetes in mice (Trammell et al.
2016c), peripheral neuropathy in mice (Trammell et al. 2016c), noise induced hearing loss in mice (Brown et al. 2014), dilated cardiomyopathy in mice (Diguet et al. 2018), mitochondrial myopathy in mice (Khan et al. 2014), liver fibrosis in mice (Pham et al. 2019), and non- alcoholic fatty liver disease in mice and humans (Zhou et al. 2016). In mice models, the ability
47
Chapter 1 Introduction
of NMN and NR to raise intracellular NAD+ levels has been reported consistently as around 2-
3 fold (de Picciotto et al. 2016; Trammell et al. 2016a; Yoshino et al. 2011). More recently, the
reduced form of NR, dihydronicotinamide riboside (NRH) (Giroud-Gerbetant et al. 2019; Yang
et al. 2019) was shown to increase NAD+ levels by up to 10-fold in cultured mammalian cells and multiple mouse tissues including liver, kidney, brain, muscle and adipose tissue (Giroud-
Gerbetant et al. 2019; Yang et al. 2019; Yang et al. 2020).
The mechanisms underlying the benefits of NAD+ precursor supplementation can be
largely attributed to the activation of the sirtuins and increased deacetylation of their protein
targets. For example, supplementation with NMN restored renal NAD+ content and SIRT1
activity in old mice, protecting against cisplatin-induced acute kidney injury through SIRT1-
mediated regulation of c-Jun N-terminal kinase (JNK) signalling (Guan et al. 2017b). Similarly,
administration of NR rescued alcohol-induced liver injuries by increasing NAD+ levels and
stimulating SIRT1-mediated deacetylation of PGC-1α, which promotes mitochondrial
biogenesis and function (Wang et al. 2018). There is mounting evidence to show that
stimulating the activity of the sirtuins and reducing the activity of other NAD+-consuming enzymes such as PARPs and CD38 protect against all major causes of disease and disability, from hearing and vision loss to cognitive and motor dysfunction as well as metabolic disorders, cardiovascular disease, infertility and immune deficiencies (Rajman, Chwalek, and Sinclair
2018). Increasing NAD+ levels through supplementation with NAD+ precursors, therefore, is a
promising therapeutic strategy to treat age-related diseases.
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Chapter 1 Introduction
1.6 NAD+ biosynthesis pathways
The biosynthesis of NAD+ from dietary sources or following supplementation with NAD+
precursors is maintained through three main pathways—the de novo synthesis pathway, the
Preiss-Handler pathway, and the Salvage pathway (Figure 1.4, below). These pathways are
largely conserved in bacteria, yeast, and humans, with a few important exceptions which will
be discussed in the following sections below.
1.6.1 The de novo synthesis pathway
In yeast and humans, synthesis of NAD+ via the de novo pathway involves metabolism of the
amino acid, L-tryptophan which enters the cell via neutral or proton-coupled amino acid transporters SLC7A5 (Bhutia, Babu, and Ganapathy 2015) and SLC36A4 (Pillai and Meredith
2011; Thwaites and Anderson 2011), respectively. Once inside, L-tryptophan is converted to
N-formyl-kynurenine by tryptophan 2,3-dioxygenase (TDO, EC 1.13.11.11) (Lewis-Ballester et al. 2016; Meng et al. 2014). N-formyl-kynurenine is an unstable intermediate and is immediately hydrolysed into L-kynurenine by kynurenine formamidase (KFA, EC 3.5.1.9)
(Pabarcus and Casida 2002). L-kynurenine is then converted into 3-hydroxykynurenine via kynurenine 3-hydroxylase (KMO, EC 1.14.13.9) which requires NADPH (Breton et al. 2000).
Next, 3-hydroxykynurenine is converted into 3-hydroxyanthranilic acid via kynureninase
(KYU, EC 3.7.1.3) (Alberati‐Giani et al. 1996). From this, the intermediate, 2-amino-3- carboxymuconic semialdehyde, is formed via 3-hydroxyanthranolic acid oxygenase (3-HAO,
EC 1.13.11.6) which then undergoes non-enzymatic spontaneous condensation to form quinolinic acid (QA) (Malherbe et al. 1994). TDO, KFA, KYU and 3-HAO are all localised in the cytosol whereas KFA, in addition to the cytosol, is also found in the nucleus, and KMO is found in the outer membrane of mitochondria. The yeast homologues for TDO, KFA, KMO,
KYU and 3-HAO are mediated by Bna proteins, Bna2 (Iwamoto et al. 1995), Bna7 (Wogulis
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Chapter 1 Introduction
et al. 2008), Bna4 (Panozzo et al. 2002), Bna5 (Panozzo et al. 2002) and Bna1 (Kucharczyk et
al. 1998), respectively, leading to production of QA.
1.6.2 The Preiss-Handler pathway
Quinolinic acid phosphoribosyltransferase (QAPRT, EC 2.4.2.19) catalyses the formation of
nicotinic acid mononucleotide (NaMN) from QA, requiring both 5-phosphoribosyl-1-
pyrophosphate (PRPP) (Fukuoka, Nyaruhucha, and Shibata 1998). NaMN may also be
synthesised from either nicotinic acid (NA) via nicotinic acid phosphoribosyltransferase
(NAPRT, EC 6.3.4.21) which requires ATP and PRPP (Galassi et al. 2012) or nicotinic acid
riboside (NaR) via nicotinamide riboside kinases (NRK1 or NRK2, EC 2.7.1.22) which also
requires ATP (Khan, Xiang, and Tong 2007; Tempel et al. 2007). NA is transported into the
cell via solute carriers SLC5A8 (Gopal et al. 2005) and SLC22A13 (Bahn et al. 2008) while
NaR may enter through equilibrative nucleoside transporters SLC29A1 and SLC29A2
(Baldwin et al. 2004).
NaMN is then converted into nicotinic acid adenine dinucleotide (NaAD) via
nicotinamide mononucleotide adenylyltransferases (NMNAT1-3, EC 2.7.7.18) in the presence
of ATP (Berger et al. 2005; Garavaglia et al. 2002; Raffaelli et al. 2002; Schweiger et al. 2001;
Zhang et al. 2003). Finally, NaAD is amidated by the glutamine-dependent enzyme, NAD+
synthetase (NADSYN1, EC 6.3.5.1) for the final synthesis of NAD+ which is also an ATP-
dependent reaction (Hara et al. 2003).
In mammals, QAPRT, NAPRT, NRK1/2, NMNAT2 and NADSYN1 are localised to
the cytosol whereas NMNAT1 is nuclear. NMNAT2, in addition to the cytosol, is also found
in the Golgi apparatus and NMNAT3 is mostly mitochondrial but also found in the cytosol.
The yeast homologues for QAPRT, NAPRT, NRK1/2, NMNAT1-3 and NADSYN1 are Bna6
(Di Luccio and Wilson 2008), Npt1 (Rajavel et al. 1998), Nrk1 (Bieganowski and Brenner
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Chapter 1 Introduction
2004), Nma1/2 (Emanuelli et al. 2003; Emanuelli et al. 1999), Qns1 (Suda et al. 2003),
respectively.
1.6.3 The Salvage/Recycling pathway
The salvage pathway is considered the main NAD+ biosynthesis pathway in mammals. It
involves the “salvage” of nicotinamide (NAM) which is generated from cleavage of NAD+
from NAD+-consuming activities. This conserves the nicotinyl ring and lowers the need for
exogenous precursors such as tryptophan and NA to maintain intracellular NAD+ levels. NAM
may also enter the cell directly from external sources such as diet or supplementation with
exogenous nicotinamide through facilitated diffusion (Schuette and Rose 1983; Sofue et al.
1991). In the presence of PRPP, NAM is converted into nicotinamide mononucleotide (NMN)
via nicotinamide phosphoribosyltransferase (NAMPT, EC 2.4.2.12), the rate-limiting enzyme in this pathway (Revollo, Grimm, and Imai 2004). Interestingly, there is no yeast homologue for NAMPT, hence utilisation of NAM in yeast involves its deamidation into NA via Pnc1
(nicotinamide deamidase, EC 3.5.1.19) (Hu et al. 2007). NA may then synthesise NAD+
through the Preiss-Handler pathway, which is considered the preferred pathway for NAD+
biosynthesis in yeast (Bedalov et al. 2003; Raj et al. 2019; Sporty et al. 2009). The yeast
homologues for NMNAT enzymes include Nma1, Nma2 and Pof1 (Emanuelli et al. 2003;
Emanuelli et al. 1999; Kato and Lin 2014).
Exogenous NMN may enter the cell directly via NMN transporter SLC12A8 (Grozio
et al. 2019a) and converted to NAD+ via NMNAT1-3, the same enzymes in the Preiss-Handler pathway used to convert NaMN to NaAD (Berger et al. 2005; Emanuelli et al. 2001; Garavaglia et al. 2002; Raffaelli et al. 2002; Zhang et al. 2003). In mammamls, NAMPT is found in all nucleus and cytoplasm but is also detected in the circulation as a form known as extracellular
NAMPT (eNAMPT) (Revollo et al. 2007). This suggests NAM may be converted into NMN via eNAMPT outside the cell, however, some studies show that other NAMPT substrates such
51
Chapter 1 Introduction
as ATP and PRPP are not available in sufficient quantities in the extracellular space for this to
take place (Hara et al. 2011), warranting further investigations.
In mammals and in yeast, the salvage pathway also involves NAD+ synthesis from
another exogenous precursor, nicotinamide riboside (NR). NR may enter directly via
equilibrative nucleoside transporters (ENT) SLC29A1 and SLC29A2. Once inside, NR may be
phosphorylated to form NMN via NRK1 or NRK2 in an ATP-dependent manner (Khan, Xiang,
and Tong 2007; Tempel et al. 2007). Alternatively, NR can be hydrolysed to form NAM and
ribose-1-phosphate via purine nucleoside phosphorylase (PNP, EC 2.4.2.1) and requires
inorganic phosphate (Ealick et al. 1990). PNP is also responsible for the conversion of NaR to
NA which may then be converted to NaMN to synthesise NAD+ via the Preiss-Handler pathway. There are three yeast homologues for mammalian PNP, called Urh1, Pnp1 and Meu1
(Belenky et al. 2007), while the yeast homologue for NRK1/2 is Nrk1.
In yeast and mammals, NR and NaR metabolism via NRK1 or NRK2 is considered more physiologically relevant than their metabolism via purine nucleoside phosphorylases
(Pnp1, Urh1, Meu1 and PNP), which may be more condition-specific (e.g. if NRK-dependent pathways are inhibited). Purine nucleoside phosphorylases have greater catalytic efficiencies
(Kcat/Km) towards other substrates such as inosine than NR and NaR (Belenky et al. 2009).
Similarly, human NRK1 and NRK2 have greater catalytic efficiency towards NR and NaR than other substrates such as cytidine (Tempel et al. 2007). Hence, compared to NR metabolism via
PNP pathways, NRK-dependent pathways are considered more physiologically relevant in mammals.
Recently, the reduced form of NR, dihydronicotinamide riboside (NRH) was reported as another precursor for NAD+ biosynthesis, which increased NAD+ up to 10-fold in cells and
tissues (Giroud-Gerbetant et al. 2019; Yang et al. 2019; Yang et al. 2020), more than any other
52
Chapter 1 Introduction
known NAD+ precursor which typically increase NAD+ levels 2 to 3-fold (Mills et al. 2016;
Trammell et al. 2016a; Yoshino et al. 2011). These authors report that NRH can be converted
to NMNH, the reduced form of NMN, via adenosine kinase (AK), where NRH is in competition
with adenosine, another substrate for AK. They further show that NMNH is then adenylated to
NADH via NMNATs which is then oxidised to form NAD+, revealing a completely new
pathway for NAD+ biosynthesis.
1.6.4 Additional pathways in yeast and bacteria, that do not exist in mammals
In yeast and bacteria (E. coli), NAM can be deamidated via Pnc1 (Hu et al. 2007) and PncA
(Frothingham et al. 1996), respectively, to form NA, a pathway which does not seem to exist in mammals (Rongvaux et al. 2003). NA may then be converted into NaMN via Npt1 in yeast
(Rajavel et al. 1998) and PncB in bacteria (Wubbolts et al. 1990) to synthesise NAD+ via the
Preiss-Handler pathway. Bacteria also possess another deamidase enzyme called NMN deamidase, PncC (Galeazzi et al. 2011), forming NaMN which can also synthesise NAD+ via
the Preiss-Handler pathway. NAD+ biosynthesis pathways in bacteria and their contribution to mammalian NAD+ metabolism will be discussed in greater detail in Chapter 4.
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Chapter 1 Introduction
Figure 1.4 NAD+ biosynthesis pathways in mammals. In mammals, NAD+ can be synthesised via three pathways. In the de novo synthesis pathway, L-tryptophan is transported into the cell via solute carriers SLC7A5 and SLC36A4 and converted into quinolinic acid (QA) via the kynurenine pathway in five enzymatic and one non-enzymatic steps. QA is then converted into nicotinic acid mononucleotide (NaMN) via quinolinic acid phosphoribosyltransferase (QAPRT) which requires 5-phosphoribosyl-1- pyrophosphate (PRPP). As part of the Preiss-Handler pathway, NaMN can also be synthesised from nicotinic acid riboside (NaR) and nicotinic acid (NA) via nicotinamide riboside kinases 1 and 2 (NRK1/2) and nicotinic acid phosphoribosyltransferase (NAPRT) respectively, which both require ATP (adenosine triphosphate). NaR is transported by equilibrative nucleoside transporters SLC29A1 and SLC29A2 while NA may enter via SLC5A8 and SLC22A13. NaMN may then be adenylated to form nicotinic acid adenine dinucleotide (NaAD) via nicotinamide mononucleotide adenylyltransferases 1, 2 and 3 (NMNAT1-3), which also requires ATP. NaAD is then amidated to nicotinamide adenine dinucleotide (NAD+) via the glutamine-dependent NAD synthetase (NADSYN1) and requires ATP. In the Salvage pathway, nicotinamide (NAM) is taken up via diffusion and converted into nicotinamide mononucleotide (NMN) via the rate-limiting enzyme nicotinamide phosphoribosyltransferase (NAMPT) and requires PRPP. NMN can also be derived from dephosphorylation to nicotinamide riboside (NR) by ecto-‘5’-nucleotidase CD73 prior to entry. NR may then enter via equilibrative nucleoside transporters SLC29A1 and SLC29A2 and re-phosphorylated into NMN via NRK1/2 which requires ATP. NR may also be degraded into NAM and ribose-1-phosphate via purine nucleoside phosphorylase (PNP) which requires inorganic phosphate. NMN may also enter directly via SLC12A8 and is then converted to NAD+ via NMNAT1-3 in an ATP-dependent manner. NAD+ is cleaved by NAD+-consuming enzymes such as the sirtuins, poly-ADP-ribose polymerases (PARPs), NAD+ glycohydrolases CD38/CD157 and sterile alpha and Toll/interleukin-1 receptor motif- containing 1 (SARM1) generating ADP-ribose moiety and free NAM as by-products. NAM is recycled back into the Salvage pathway to synthesise more NAD+ and maintain intracellular NAD+ levels without being limited by the uptake of extracellular precursors.
54
Chapter 1 Introduction
1.7 Current gaps in the field of NAD+ biology
Despite extensive research in the field of NAD+ biology and its implications in health and
disease, there are major questions that are yet to be explored regarding NAD+ precursor metabolism.
1.7.1 Unexplored mechanisms of NAD+ precursor deamidation
Given the interest in using NAD+ precursors to improve healthspan and treat disease, it is
important to understand the metabolism of NAD+ precursors in mammalian systems and the pathways involved in their integration into the NAD+ metabolome. As described in Section
2.6.4 (above), NAM may be deamidated to form NA by Pnc1 in yeast and PncA in bacteria,
while NMN can be deamidated to form NaMN by PncC in bacteria. Mammals are not known
to have an sequivalent step, though some evidence suggests that deamidation of NR in
mammalian systems may exist.
Trammell and colleagues observed the appearance of the deamidated metabolite NaAD
in mouse liver and human peripheral blood mononuclear cells (PBMCs) following oral
administration of NR, an amidated NAD+ precursor. This was unexpected, as there is no
pathway in mammals by which NR can be integrated into NaAD. As part of the Preiss-Handler
pathway, NaAD is amidated to form NAD+ via the glutamine dependent enzyme NAD
synthetase (NADSYN1) (Hara et al. 2003), while NR is phosphorylated into NMN by the enzymes NRK1/2 (Tempel et al. 2007). One possible explanation for NR-derived NaAD formation could be deamidation of NMN to NaMN, the precursor to NaAD in the Preiss-
Handler pathway. Importantly, as there are no mammalian homologues of the bacterial NMN deamidase PncC, orally administered NMN may be deamidated by bacterial PncC in the gut.
This hypothesis, which is investigated in Chapter 4 and Chapter 5 of this thesis, is an important step in understanding the contribution of the microbiome to the metabolism of orally administered NAD+ precursors which may affect their therapeutic efficacy. Further, these 55
Chapter 1 Introduction
investigations may be important for the therapeutic development of NAD+ precursors, in terms
of dosing and routes of administration.
1.7.2 NAD+ precursor uptake mechanisms still unclear
With the exception of studies in neurons (Ying et al. 2007; Zheng et al. 2012) and
cardiomyocytes (Liu et al. 2014; Zhang et al. 2016b), NAD+ is thought to be unable to cross
the plasma membrane, due to its size (663.43 g/mol) and highly polar structure (Di Lisa and
Ziegler 2001). While an NAD+ transporter has been identified in bacteria (Haferkamp et al.
2004) as well as in the mitochondria of yeast (Todisco et al. 2006) and plants (Palmieri et al.
2009), it is unclear whether one exists in mammals. Dedicated NAD+ transporters are thought
to exist in mammalian cells as connexin 43 channels (Bruzzone et al. 2001) however, they may only be expressed in certain cell types (Billington et al. 2008; Pittelli et al. 2011; Wang et al.
2008; Ying et al. 2007) and supplementation with NAD+ precursors is more effective at raising intracellular NAD+ levels across different cell types and tissues.
In mammalian cells, tryptophan uptake occurs via solute carrier proteins that also
transport large uncharged amino acids (Palego et al. 2016). NA is also taken up by specific,
high affinity solute carrier proteins that involve an acidic pH, temperature and energy-
dependent anion antiporter or a proton cotransporter (Nabokina, Kashyap, and Said 2005; Said
et al. 2007; Takanaga et al. 1996). The exact mechanism of NAM is not completely understood,
but is considered to involve facilitated diffusion (Schuette and Rose 1983; Sofue et al. 1991) or conversion into NMN, as extracellular NAMPT is also present in the circulation (Revollo et al. 2007), though some evidence suggests that other NAMPT substrates such as ATP and PRPP are not available in sufficient quantities in the extracellular space (Hara et al. 2011). In the latter case, transport of NMN into the cell remains a controversial matter in the field of NAD+
biology and will be investigated in Chapter 5 of this thesis. Briefly, the debate is around
whether the uptake of exogenously supplemented NMN occurs directly, through transporter
56
Chapter 1 Introduction
SLC12A8 (Grozio et al. 2019a) or first requires dephosphorylation into its nucleoside form,
NR (Ratajczak et al. 2016). NR and NaR may be directly transported via equilibrative
nucleoside transporters (ENTs) SLC29A1 and SLC29A2 (Mangravite, Xiao, and Giacomini
2003) and re-phosphorylated into NMN and NaMN respectively, via NRK1 or NRK2
intracellularly (Sasiak and Saunders 1996; Tempel et al. 2007). While separate studies have
shown NMN and NR are both effective at raising intracellular NAD+ levels up to 2 to 3-fold
(Trammell et al. 2016a; Yoshino et al. 2011), understanding how these precursors are
transported and integrated into the NAD metabolome could improve their therapeutic potential.
1.7.3 Quantifying the NAD+ metabolome remains a challenge
Given the important role of NAD+ in health and disease, accurate measurement of NAD+ and
its metabolites is essential. These have been measured using a variety of methods. NAD+ may
be measured using enzymatic and colorimetric assays (Bernofsky and Swan 1973; Zhu and
Rand 2012), though these methods are often indirect, cannot detect low picomolar levels (thus
requiring more sample), and are easily influenced by minor changes in temperature and pH,
hence difficult to reproduce. To increase sensitivity, reverse-phase high-performance liquid
chromatography (HPLC) methods that use mobile phases with buffer salts and ion pairing
agents have been employed however, detecting changes in low picomolar ranges in complex
biological samples such as cell lysates and tissue homogenates remains a challenge (Casabona
et al. 1995). These techniques are also limited by their low throughput, measuring only one or
two metabolites at one time, and are unable to accurately separate metabolites that are similar
in biochemical properties and molecular structure in complex biological samples.
The ideal method to quantify the NAD metabolome must be rapid, reproducible, and
robust. Currently, the leading technique to quantify NAD+ metabolites is liquid chromatography coupled to tandem mass spectrometry (LC-MS/MS). Its greatest advantage is being able to detect multiple metabolites in a single injection, which can be aided by using
57
Chapter 1 Introduction multiple reaction monitoring (MRM) mode to identify metabolites of interest as a function of time, which also increases the specificity of measurements. One major challenge, however, is the lack of standardised methods to extract, detect and analyse metabolites of interest, often requiring time-consuming optimisation to accurately measure metabolites of interest with good resolution while minimising signal-to-noise ratios. Separating metabolites with similar biochemical properties (e.g. mass, polarity, charge), also remains as a major challenge in quantification of the NAD metabolome as there are many closely related pairs of metabolites such as the amide/acid pairs, NAM/NA, NMN/NaMN, NAD+/NaAD, and NR/NaR, which all differ from each other by one Dalton. With more studies seeking to measure NAD+ and its metabolites in health and disease, these challenges need to be overcome and an improvement in methods to quantify the NAD+ metabolome is warranted.
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Chapter 1 Introduction
1.8 Thesis goals
This thesis aims to investigate gaps in knowledge in the field of NAD+ precursor metabolism.
These studies focused on the metabolism of NMN as an emerging NAD+ precursor with
promising therapeutic potential, which has already started to undergo Phase I clinical trials in
humans (Irie et al. 2019, 2020). Important to the clinical translation of NMN, is an accurate
understanding of its metabolism through the oral route, which is the most common route of
administration in the clinical setting.
To explore the pathways involved in metabolism of orally administered NMN, we
strategically designed labelled isotopes of NMN at the ribose (13C) and nicotinamide moieties
(15N) (Chapter 2 Section 2.1) and developed an LC-MS/MS method to trace the incorporation of heavy labels into NAD+ metabolites (Chapter 3). This method was validated in mammalian cell lines to verify our labelling strategy and ensure utilisation of NMN occurred via the expected canonical routes. The overarching hypothesis was that the microbiome contributes to host NAD+ metabolism by metabolising orally delivered NMN via the deamidated route, giving
rise to the deamidated precursors NaMN and NaAD. To investigate this hypothesis, this project
first explored NMN utilisation in microbial systems, supplementing NMN into the culture broth
of E. coli bacteria and measuring changes to NAD+ metabolites in supernatants and cell lysates
(Chapter 4). As expected, deamidation of NMN via PncC was observed, measured by increases
in levels of NaMN in cell lysates and allowing us to explore our hypothesis further in vivo
(Chapter 5).
To investigate the contribution of the microbiome to the metabolism of NMN, labelled
NMN isotopes were orally delivered by gavage to antibiotic-treated mice, to ablate the
microbiome, and changes in the labelled NAD metabolome was detected in the intestinal
tissues, liver and plasma. These findings suggest the microbiome is in competition with the
host for orally administered NAD+ precursors such as NMN, consuming NAD+ metabolites for 59
Chapter 1 Introduction
their own growth and survival and making them less available to the host. Further, while there
was very little evidence to support the deamidation of NMN to NaMN, important factors such
as sampling at a single time point may have influenced these results. Interestingly, an
abundance of NR was observed following supplementation with NMN both in vitro and in vivo,
supporting the indirect mechanism of NMN transport which involves dephosphorylation to NR.
Overall, the findings from this thesis have important implications in the dosing requirements
and route of administration of NAD+ precursors such as NMN and NR, where their co- administration with antibiotics may improve therapeutic efficacy to delay ageing and treat age-
related diseases.
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Chapter 2 General Methods
General Methods
2.1 Isotopic labelled compounds
2.1.1 Synthesis of labelled NMN compounds
Synthesis of our labelled NMN compounds employed a two-step process; 1) the initial synthesis of isotope labelled nicotinamide and ribose, followed by 2) an enzyme-mediated ligation of these isotope labelled constituents into NMN. The ribose moiety was 13C labelled
at all five carbon positions (purchased from Cambridge Isotope Laboratories, Cat. No. CLM-
3652) and was used to synthesise both NMN compounds. Nicotinamide was 15N labelled at either the base nitrogen of the nicotinyl ring or at both the base and the amide, following custom synthesis by Dr. Sebastian Marcuccio, Advanced Molecular Technologies, VIC. To combine the labelled ribose and labelled nicotinamide moieties an enzyme-based reaction was fuelled by two enzymes, recombinant phosphoribosyl synthetase (PRS) and recombinant nicotinamide
61
Chapter 2 General Methods phosphoribosyltransferase (NAMPT) (Wu, Sinclair, and Meetze 2019). NAMPT and PRS were added into a reaction buffer containing 1mM labelled ribose, 1mM labelled nicotinamide, 3mM
ATP, 1mM dithiothreitol, 1 mM MgCl2 and 50mM Tris-HCl (pH 7.5) and incubated at 37°C for 30 minutes. The reaction was terminated with the addition of 0.01% trichloroacetic acid
(TCA). The final compounds were name NMN1 and NMN2. NMN1 refers to the NMN isotope labelled at the ribose and base nitrogen of the nicotinyl ring (hereafter referred to as base) with a final mass shift of M+6 (MW: 340.2g/mol) (Figure 2.1 A) and NMN2 refers to the NMN isotope labelled at the ribose and at both the base and amide nitrogen positions with a final mass shift of M+7 (MW: 341.2g/mol) (Figure 2.1 B). Labelled isotopes were purified with size-exclusion and ion exchange columns and concentrated by lyophilization to yield >95% purity. Labelling was confirmed by mass spectrometry (Figure 2.2). The stability of labelled
NMN compounds were not tested, however, were stored either as powder or aliquoted at a concentration of 20mM, both preparations kept at -30℃.
Figure 2.1 Chemical structure and isotope labelling positions of NMN (A) NMN1 (M+6) labelled with five 13C at the ribose moiety and with 15N at the pyridine base of the nicotinamide moiety. (B) NMN2 (M+7) labelled with five 13C at the ribose moiety and with 15N at the pyridine base and amide of the nicotinamide moiety.
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Chapter 2 General Methods
Figure 2.2 Chromatogram peaks of NMN isotopes using LC-MS/MS injected at 100 µM. (A) Unlabelled NMN (M+0) with MRM transition 335.15123.05. (B) Labelled NMN1 (M+6) with MRM transition 341.15124.05. (C) Labelled NMN2 (M+7) with MRM transition 342.1500125.05.
2.1.2 Labelled glutamine
15N-amide-glutamine (>98%) was purchased from Cambridge Isotope Laboratories (cat.no.
NLM-557-0) (Figure 2.3) and stored as powder at room temperature or in aliquots at a stock concentration of 200 mM at -30℃. Chromatograms of labelled glutamine peaks were confirmed by mass spectrometry and are shown in Figure 2.4.
Figure 2.3 Chemical structure and isotope labelling of glutamine.
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Chapter 2 General Methods
Figure 2.4 Chromatogram peaks of unlabelled glutamine and isotope labelled glutamine using LC-MS/MS injected at 100 µM. (A) Unlabelled glutamine (M+0) with MRM transition 147.1444. (B) 15N-amide-labelled glutamine (M+1) with MRM transition 148.1445.
2.2 Sample processing for mass spectrometry
2.2.1 Preparation of extraction buffer
The extraction buffer for cell and tissue processing was acetonitrile (Ajax Finechem, cat.no.
2315), methanol (Ajax Finechem, cat.no. 2314) and deionized water (Merck MilliQ®) at a 2:2:1
(v/v) ratio. A mixture of the internal standards, methionine sulfonate (MES), camphorsulfonic acid (CSA) and thymine-d4 (TD4) was diluted into the acetonitrile:methanol:water mixture from a 200X concentrated stock to obtain a 0.1X concentration (targeting in-vial final concentration of 0.25 µM CSA, 5 µm MES and 5 µM TD4). The solution was pre-cooled to -
30℃ overnight before use (the freezing point of acetonitrile and methanol is -45℃ and -97.6℃ respectively) and maintained on ice during sample extraction.
64
Chapter 2 General Methods
2.2.2 Cell harvest and extraction
The following cell harvest protocol was applied to primary hepatocytes, C2C12 and HEK293
cells (see Chapter 3 for cell culture conditions). For cell harvest, media was removed before
rinsing the cells one with phosphate buffered saline (PBS) cooled to 4°C. Cells were extracted
based on a method previously published by Yuan and colleagues (Yuan et al. 2012). Five
hundred microliters of cold acetonitrile/methanol/water extraction buffer, as above, was added.
Using a cell scraper, the cells in extraction buffer were transferred into microcentrifuge tubes
and centrifuged at 16,000g for 10 minutes at 4°C. The supernatant was transferred into a clean
microcentrifuge tube and snap frozen immediately in liquid nitrogen for storage at -80°C until
further processing. The remaining pellet was used to determine protein concentration (Pierce™
BCA Protein Assay kit, Thermo Fisher Scientific, cat.no. 23225) for normalisation of data
during data analysis.
2.2.3 Further sample processing for mass spectrometry
All samples were thawed on ice and centrifuged briefly before being dried using a speed
vacuum system (Savant SpeedVac SPD140DDA Vacuum Concentrator, Thermo Fisher
Scientific) for approximately 2-3 hours at vacuum. The dried pellet was resuspended in 50 µL
of LC-MS-grade water (Fisher Healthcare, cat.no. PI51140) and centrifuged at 16,000g for 10 minutes at 4°C. The supernatant was transferred into HPLC vials and analysed promptly by
LC-MS/MS.
2.2.4 NAD+ metabolite standard curve preparation
Primary stock solutions of NAD+ metabolite standards were stored at -30℃ at concentrations
of 1 mM and above. Working standard solutions were stored at -30℃ at 100µM concentrations
in 100 µL aliquots to avoid multiple freeze-thaws. The standard calibration series of NAD+
metabolites were generated by performing two-fold serial dilutions of the 100 µM standard
65
Chapter 2 General Methods stock solution in HPLC-grade water. Standard calibration curves were generated using LC-
MS/MS for the following NAD+ metabolites; NAD+ (Sigma-Aldrich, cat.no. N7004), NAM
(Sigma-Aldrich, cat.no. N3376), NMN (GeneHarbour Biotechnologies, Hong Kong), NR
(synthesized by Dr. Hamish Toop, School of Chemistry, UNSW), NaR (synthesized by Dr.
Hamish Toop), NA (Sigma-Aldrich, cat.no. 72309), NaAD (Sigma-Aldrich, cat.no. N4256), and NaMN (GeneHarbour Biotechnologies, Hong Kong). The standard calibration series was prepared on the same day as further processing of samples for LC-MS/MS analysis.
2.3 LC-MS/MS
2.3.1 Liquid chromatography
High performance liquid chromatography (HPLC) was performed using the 1260 Infinity LC
System (Agilent) with a chilled autosampler set at 4-8℃. LC separation was accomplished on an amide capped ethylene-bridged hybrid (Amide XBridge BEH) HPLC column (100 mm x
2.1 mm, 3.5 µm particle size, Waters Corporation) at room temperature (Yuan et al. 2012). For the mobile phase, Buffer A, consisted of a 95:5% (v/v) mixture of water and acetonitrile
(CH3CN) containing 20 mM ammonium acetate (NH4CH3CO2) and 20mM acetic acid
(CH3COOH) at a final pH 5 which was stored at room temperature for up to 2 weeks. This acidic buffer was adapted based on the formulation described in Yuan et al. (2012), which contained 20mM ammonium hydroxide instead of acetic acid, and is a formulation that is commonly used (Anastasiou et al. 2011; Asara et al. 2009; Caroline et al. 2011; Gao et al. 2011;
Kelly et al. 2011; Locasale et al. 2011; Locasale et al. 2012; Vander Heiden et al. 2010). In this study, however, metabolites with the same MRM transitions eluted at similar retention times causing an overlap of peaks that were unable to be resolved during peak integration. In order to achieve separation of these peaks, the pH of Buffer A was lowered from 9.4 to 5 by replacing
20mM ammonium hydroxide with 20mM acetic acid. The acidic nature of the buffer interacts with the hydroxyl group from acidic NAD+ metabolites including NaMN, NaAD, NA and NaR,
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Chapter 2 General Methods
separating them from NMN, NAD+, NAM and NR, respectively, to achieve better integration
of peaks. Buffer B was 100% acetonitrile (CH3CN) and was stored for up to 6 months at room
temperature. The flow rate was 200 µl per minute, and the buffers were flowed in gradient
mode with percentages of Buffer B set at 85% (0 min), 85% (0.1 min), 70% (10 min), 30% (13
min), 30% (17 min), 85% (17.5 min), and 85% (30 min). The injection volume taken from each
vial was 2.5μL.
2.3.2 Mass spectrometry
HPLC was coupled to the QTRAP® 5500 mass spectrometer controlled by Analyst® Software
(AB SCIEX, Toronto, Canada). The following MS settings were applied for MS/MS detection
of metabolites of interest: the ion source was set at 350°C and ionisation voltage at 4500 volts
with positive/negative polarity switching. Mass isotopologues of metabolites were acquired by
MS2, using the unscheduled multiple reaction monitoring (MRM) mode with a dwell time of
40 ms. The MS parameters consisting of the declustering potential (DP), collision energy (CE) and cell exit potential (CXP), and MRM transitions were optimised based on the monoisotopic mass of chemical standards.
2.4 Data analysis
For experiments using labelled compounds, MSConvert (version 3.0.18165-fd93202f5) and in-
house MATLAB scripts were employed by co-supervisor Dr. Lake-Ee Quek who kindly integrated the metabolite peaks and extracted the data. Deconvolution scripts were developed to resolve the overlapping pairs of closely related metabolites as in Table 3.2: NaAD
(665.03>428.06) & NAD+ (664.0>428.09), NaR (256.2>124.2) & NR (255.0>123.03) and
NaMN (336.10>123.78) & NMN (335.15>123.05) using MATLAB’s Optimisation Toolbox™
(version R2018a, The MathWorks, Natick, MA, USA). The integrated peak areas were
normalised first, by dividing by the integrated peak areas of the internal standards, to account
67
Chapter 2 General Methods for any sample loss during preparation, and then normalised again by dividing by the amount of protein (for cells), optical density values (for E.coli) or frozen tissue weight (for mouse tissue).
2.5 Statistics
All data are represented as mean ± s.d. and made using GraphPad Prism 8 software (version
8.3.0). Statistical analysis tests included testing for outliers using the ROUT method with
ROUT coefficient Q set at 1% and two-way ANOVA with Sidak’s multiple comparisons test to compare whether there were differences between two or more treatment groups. A p-value less than 0.05 was considered statistically significant.
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Chapter 3 Quantifying the NAD metabolome
Development of an LC-MS/MS method to quantify the labelled NAD+ metabolome
3.1 Introduction
There is mounting evidence for the importance of NAD+ levels in health and disease. This
abundant cofactor is vital for maintaining redox homeostasis and plays an important role as a
substrate for enzymes involved in processes of cell survival including energy metabolism,
DNA damage repair, gene expression and cell signalling in response to cellular stress (Braidy
et al. 2018; Canto, Menzies, and Auwerx 2015; Chini, Tarragó, and Chini 2017; Connell,
Houtkooper, and Schrauwen 2019; Demarest et al. 2019; Johnson and Imai 2018; Kulkarni and
Brookes 2019). As NAD+ reportedly declines with age and in various pathological conditions
(Camacho-Pereira et al. 2016; Clement et al. 2019; Das et al. 2018; Frederick et al. 2016;
Gomes et al. 2013; Massudi et al. 2012b; Schultz and Sinclair 2016), the use of NAD+
precursors such as NMN and NR are being pursued to raise NAD+ levels to restore the cell
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Chapter 3 Quantifying the NAD metabolome back into a youthful and healthy state (Caton et al. 2011; Chambon, Weill, and Mandel 1963; de Picciotto et al. 2016; Grozio et al. 2019a; Lu et al. 2014a; Mills et al. 2016; Park et al. 2016;
Rajman, Chwalek, and Sinclair 2018; Tarantini et al. 2019; Yamamoto et al. 2014b; Yao et al.
2017; Yoshino et al. 2011; Zhang et al. 2017).
The efficacy of NAD+ precursor supplementation is largely assessed by measurement of NAD+ itself, and less frequently, its metabolites. The therapeutic benefits of NMN are largely attributed to an increase in intracellular NAD+, however the precise mechanisms of
NMN transport and utilisation are controversial. Unanswered questions include the effect of
NMN on the overall NAD+ metabolome, including its impact on other metabolites. Other
NAD+ precursors such as nicotinic acid (NA) and nicotinamide (NAM) (collectively termed niacin) as well as nicotinamide riboside (NR) have been shown to increase levels of metabolites in the Preiss-Handler pathway such as nicotinic acid adenine dinucleotide (NaAD) and nicotinic acid mononucleotide (NaMN) in mouse studies (Trammell et al. 2016a). NA is a deamidated metabolite in the Preiss-Handler pathway, which can be converted to NaMN via
NA phosphoribosyltransferase (NAPRT) (Niedel and Dietrich 1973; Preiss and Handler
1958b). NaMN is then converted to NaAD via NMN adenylyltransferase (NMNAT) (Berger et al. 2005; Raffaelli et al. 2002; Schweiger et al. 2001; Zhang et al. 2003) before NaAD is amidated via the glutamine-dependent NAD+ synthetase (NADSYN1) in the final step of
NAD+ synthesis (Hara et al. 2003). As expected, NA treatment increases the formation of the deamidated metabolites NaMN and NaAD (Preiss and Handler 1958a). Surprisingly, these deamidated metabolites are also increased following supplementation with the amidated precursors NAM and NR, which are metabolised in the NAD+ salvage pathway (Trammell et al. 2016a). Evidence for the existence of deamidase enzymes in mammals is limited, with papers investigating this dating 4-5 decades ago failing to identify a mammalian enzyme that could mediate this activity, despite trace levels of nicotinamide deamidation being detected
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(Petrack et al. 1963; Petrack et al. 1965; Sarma, Rajalakshmi, and Sarma 1961; Tanigawa et al.
1971). Given recent findings reporting the appearance of deamidated metabolites following
administration with the amidated precursor NR, this pathway is worthy of further investigation.
Isotopic labelling studies allow for tracing the metabolic fate of compounds, which
could be used here to may enhance our understanding of how various NAD precursors are
incorporated into the NAD metabolome. After the initial report by Trammel and colleagues,
recent studies have used labelled versions of NMN to investigate the mechanisms of transport,
utilisation and bioavailability based on different routes of administration in mice including oral
gavage and intraperitoneal injections (Grozio et al. 2019a; Liu et al. 2018; Ratajczak et al.
2016). In these studies, the incorporation of labelled ribose and nicotinamide moieties from
NMN into NAD+ was traced, but their incorporation into other members of the NAD+
metabolome such as deamidated metabolites, NaMN and NaAD was lacking. The ability to
trace labelled ribose and nicotinamide moieties from isotope labelled versions of NAD+
precursors as they are incorporated into NAD+ metabolites is an important opportunity to
investigate the pathways involved in NMN uptake and utilisation, and its impact on other metabolites contributing to NAD+ biosynthesis in mammalian cells.
Liquid chromatography coupled to tandem mass spectrometry (LC-MS/MS) is the
gold-standard for the reliable analysis of NAD+ metabolites in mammalian cells (Bustamante
et al. 2018; Liang et al. 2014; Martino Carpi et al. 2018; Trammell and Brenner 2013; Yaku,
Okabe, and Nakagawa 2018b). In a typical mass spectrometer, quadrupoles made up of four
parallel metal rods allow the transmission of a voltage to be applied along their axis, to act as
mass filters for ionised compounds. Tandem mass spectrometers comprise two quadrupoles
flanking a collision cell, a third quadrupole, that enable mass spectrometry to be performed one
after another (MS/MS). The samples are first ionised through a process called electrospray
ionisation (ESI) (Bruins 1998; Ho et al. 2003) which gives each compound an additional
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charge. As these ions move through the electrical and magnetic fields of voltage-applied
quadrupoles, their movement is affected such that they may be selectively filtered based on
their mass to charge ratio (m/z). The first quadrupole selects the complete target metabolite,
called the precursor ion (or Q1). It then enters the collision cell (second quadrupole) where it
is fragmented into smaller ions using an inert gas, such as nitrogen or argon, in a process called
collision induced dissociation (CID) (Wells and McLuckey 2005). These small ions, called
product ions (or Q3) then enter the third quadrupole for selection and detection to generate a
mass spectrum with chromatogram peaks whose area can be quantified and analysed for a
metabolite of interest. This process is called selective reaction monitoring (SRM), which when
applied simultaneously to select and detect multiple precursor and product ion pairs, is called
multiple reaction monitoring (MRM).
An important aspect of using MRM is identifying the MRM transition value for each
metabolite, which refers to the m/z values of each precursor/product ion pair as a unique
identifier of each metabolite. This increases the specificity and the capacity to multiplex for
multiple metabolites which results in the high-throughput nature of this technique. In targeting
NAD+ metabolites in this study, the mass to charge (m/z) for each parent ion was based on the
monoisotopic mass of each compound derived from PubChem identity searches
(https://pubchem.ncbi.nlm.nih.gov). Nicotinamide adenine dinucleotide (NAD+), nicotinamide riboside (NR) and nicotinic acid adenine dinucleotide (NaAD) are the only NAD+ metabolites
from our metabolites of interest with a formal positive charge. As such, there is no change in
the parent ion m/z ratio and electrospray ionisation predominantly functions to produce gas phase ions from the solution phase ions to pass it through the quadrupoles. All other NAD+
metabolites analysed, namely, NMN, NaMN, NaR, NAM and NA carry a formal charge of
zero, and therefore the ESI adds an additional charge to their monoisotopic mass as a unique
identifier of their respective MRM transitions. In order to optimise the measurement of each
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metabolite, it is necessary to optimise a series of mass spectrometry parameters including
depolarisation potential (DP), collision energy (CE), and cell exit potential (CXP).
The retention time (RT), defined by the time from injection to detection of a metabolite,
adds another level of specificity to target each metabolite of interest and is another parameter
that needs to be determined during the optimisation process. Once established, this list serves
as a set of instructions for the mass spectrometer to identify and detect metabolites of interest
in experimental samples to generate chromatogram peaks and mass spectral data. As these
parameters are non-transferable and vary significantly between different mass spectrometers, buffers and LC columns, the optimisation of methods between studies is warranted.
The first aim of this study was to develop an optimised LC-MS/MS method to measure unlabelled and labelled NAD+ isotopologues (molecules that have the same chemical formula
and structural arrangement but differ by incorporation of isotope labels). These would be
measured following supplementation with two stable isotope labelled versions of NMN, which
are designated here as NMN1 and NMN2. NMN1 was isotopically labelled at five carbons
(13C) of the ribose moiety (M+5) and at the nicotinyl ring base (15N) of the nicotinamide moiety
(M+1), providing a total mass shift of M+6 (Figure 3.1 B, below). NMN2 was labelled at the same positions as NMN1 with an additional label at the amine functional group (15N) of the nicotinamide moiety (M+2), providing a total mass shift of M+7 (Figure 3.1 C, below). These
isotopically labelled versions of NMN were strategically designed to trace the incorporation of
labelled moieties in NAD+ metabolites to investigate the metabolic fate of NMN and explore
the mechanisms of NMN transport. Following optimisation of LC-MS/MS parameters to
analyse the NAD+ metabolome, we sought to validate these measurements and trace these
isotope-labelled metabolites using well-characterised cell lines, including primary hepatocytes,
C2C12 myoblasts and HEK293 cells.
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Figure 3.1 Labelling strategy of labelled NMN isotopes. The biochemical structure and molecular weight of (A) unlabelled (M+0), (B) (M+6) labelled NMN1 or (C) (M+7) labelled NMN2 is shown above. The ribose moiety was labelled at five positions with 13C and the nicotinamide moiety was labelled either at the base or base and amide position with 15N. The ribose and nicotinamide moieties were synthesised separately before being combined to synthesise the labelled NMN compounds through an enzyme-based reaction (Wu, Sinclair, and Meetze 2019) (see Chapter 2, Section 2.1.1 for more details on the enzymatic synthesis of labelled NMN compounds).
When validating our measurement system using these cell lines, these experiments
could address a third aim of this investigation regarding the mechanism by which NMN is
transported into cells, currently a topic of debate in the field of NAD+ biology. Understanding
this mechanism could provide important insights into how cells access extracellular NMN and
in turn, assess its effectiveness as an NAD+ precursor compared to other precursors such as
nicotinamide riboside (NR).
The first described mechanism for NMN transport into cells was demonstrated by its
dephosphorylation to NR outside the cell via the ecto-5’-nucleotidase CD73 (cluster of
differentiation number 73) (Grozio et al. 2013). NR could then be taken up into the cell via equilibrative nucleoside transporters (ENTs) (Nikiforov et al. 2011; Sociali et al. 2016), to be re-phosphorylated back into NMN via NR kinases (NRK1/2) (Bieganowski and Brenner 2004;
Ratajczak et al. 2016). While it was speculated that an NMN transporter may potentially exist, a recent study described a role for the transmembrane protein SLC12A8 as an NMN transporter. This transporter belongs to the SLC12 gene family of electroneutral cation-chloride
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cotransporters, with high levels of expression in the small intestine and pancreas, and moderate
expression in the liver and white adipose tissue (Grozio et al. 2019).
Studies conducted by Ratajzcak and colleagues hypothesised that if primary
hepatocytes take up intact NMN as a substrate for NAD+ synthesis, exposure to double labelled
NMN (2H-13C-NMN), labelled at the nicotinamide (13C) and ribose (2H) moieties, would
produce both double-labelled intracellular NMN and NAD+ (Ratajzcak et al. 2016).
Interestingly, neither doubly labelled NMN or NAD+ were detected in cells following
supplementation with double labelled NMN, unless double labelled NR was used instead. This
indicated NMN dephosphorylation to NR is a key step when cells assimilate NMN. The
detection of double labelled NR following supplementation with double labelled NMN was not
reported, which could have provided further evidence to support these claims. This was,
however, measured in a recent study by Grozio and colleagues to investigate NMN transport
via the putative transporter SLC12A8, by using NMN labelled at the nicotinamide moiety (18O-
D-NMN). In contrast to the study conducted by Ratajzcak and colleagues, Grozio and
colleagues provided evidence for intact NMN transport by detecting 18O-D-NMN from primary
hepatocytes isolated from wild-type mice but not from primary hepatocytes isolated from
SLC12A8 knock out mice (Grozio et al. 2019). The absence of 18O-D-NR also led researchers
to rule out the possibility of NMN transport via dephosphorylation into NR to further support
their claims for NMN transport via SLC12A8. The short harvest time point used in this study
(5 minutes) however, may not have been long enough to observe whether dephosphorylation of NMN to NR may occur, and further studies investigating this at later time points are warranted.
Another important consideration regarding the administration of NAD+ precursors is
whether there are physiological differences in their bioavailability based on their route of
administration. The bioavailability of orally versus intravenously delivered NMN was
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investigated by Liu and colleagues, and its subsequent ability to raise NAD+ in various tissues
was measured by administering double labelled NMN (2H-13C-NMN) in mice (Liu et al. 2018).
Interestingly, intravenous delivery of labelled NMN led to an increase in labelled NAD+ in the liver and kidney, however this was absent when delivered by oral gavage, suggesting the gastrointestinal tract may play a significant role in NMN metabolism and bioavailability in mammals. The role of the gut microbiome on the metabolism of labelled NMN isotopes will be explored in subsequent chapters, Chapter 4 (exploration of microbial NMN metabolism using E.coli) and Chapter 5 (oral administration of labelled NMN in mice), while the metabolic fate of NMN and its transport into the cell using labelled NMN isotopes in in vitro models will be explored in this chapter.
The specific aims of this study were to:
1. Develop a reliable and robust LC-MS/MS method to detect the incorporation of labelled
moieties in NAD+ metabolites.
2. Trace the incorporation of NMN into the NAD+ metabolome in well-characterised in
vitro models.
3. Investigate the mechanism for NMN transport into cells by tracing the incorporation of
labelled moieties into NMN and NR.
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3.2 Methods
3.2.1 Optimisation of mass spectrometry parameters
Adopting a targeted approach, NAD+ metabolites were measured using multiple reaction monitoring (MRM), where the mass spectrometer is empirically tuned, using chemical standards, to measure an analyte of a desired mass to charge ratio (m/z) with maximum selectivity and sensitivity. The Analyst® Software (AB SCIEX, Toronto, Canada), which controls the QTRAP® 5500 mass spectrometer, can automatically select for the best MRM, i.e., the m/z transition for a pair of precursor/parent and product/fragment ion, and correspondingly, the voltages for mass spectrometry parameters, declustering potential (DP), collision energy
(CE) and collision cell exit potential (CXP). These MS parameters are analyte dependent; optimum DP, CE and CXP are required to maximise the generation of a precursor ion with the expected m/z (DP), its subsequent conversion (CE) into product ion and enrichment (CXP).
Together, these MS parameters maximise the signal of a given MRM transition.
To optimise each NAD+ metabolite, pure metabolite at 1 µM was infused at a rate of
10 µL per minute into the ion source using a syringe pump (1 mL). The Analyst® software begins the optimisation once the monoisotopic mass and number of charges for the metabolite of interest are entered. Optimisations were repeated 3 times and the average value of each parameter was calculated.
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3.2.2 Liquid chromatography (LC) separation
LC (see Section 2.3.1) was coupled to tandem mass spectrometry (MS/MS) (see Section 2.3.2)
to quantify NAD+ metabolites using the optimised MS parameters (CE, DP and CXP). The LC separation gradient is presented in Table 3.1 with a flow rate set at 200 µL per minute and the percentage of each buffer shown over the course of 30 minutes.
Table 3.1 Liquid chromatography (LC) separation gradient Time (mins) Flow rate (µl/min) Buffer A (%) Buffer B (%) 0 200 15 85 0.1 200 15 85 10 200 30 70 13 200 85 30 17 200 85 30 17.5 200 15 85 30 200 15 85
Buffer A: 95:5 (v/v) HPLC H2O:Acetonitrile (CH3CN) with 20 mM ammonium acetate
(NH4OAc) + 20 mM acetic acid (CH3COOH), pH 5. Buffer B: 100% Acetonitrile (CH3CN).
One of the challenges faced during LC was the ability to separate peaks of closely
related metabolites such as NaMN (336 m/z) from NMN (335 m/z), and NaAD (665 m/z) from
NAD+ (664 m/z). To overcome this challenge without changing the LC column, the pH of the
mobile phase Buffer A was decreased from pH 9.4, as originally described in Yuan and
colleagues (Yuan et al. 2012), to pH 5 by replacing 20 mM ammonium hydroxide with 20 mM
acetic acid. In LC-MS, adjusting the pH of the mobile phase buffers can be used as a powerful
tool to change the ionisation state of ionisable metabolites and, in turn, influence their retention
times. NA, NaR, NaMN and NaAD are all acidic metabolites characterised by a carboxylic
acid functional group which is strongly ionised at neutral pH. Under low pH conditions,
interactions between the acid buffer and the carboxylic acid will create deionised (or more
neutral) species with greater retention that ultimately allow better chromatographic separation
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with their amidated counterparts. Note the original (basic) Buffer A system did not lead to peak
separation of the acid-amide complements despite changing the LC programme/gradient.
3.2.3 Isolation of primary hepatocytes and treatment
The isolation of primary hepatocytes was based on previous work by Montgomery and
colleagues (Montgomery et al. 2015) which is described here.
Before perfusion, Hanks Buffered Salt Solution (HBSS, 1X) containing 138mM sodium chloride (NaCl), 50mM HEPES, 5.6mM glucose, 5.4mM potassium chloride (KCl),
0.34mM disodium phosphate (Na2HPO4), 0.44mM monopotassium phosphate (KH2PO4), and
® 4.17mM sodium bicarbonate (NaHCO3) was prepared in 1L of deionised water (MilliQ ,
Merck) and pH adjusted to 7.4. This solution was sterile filtered and 0.5mM ethylene glycol-
bis(β-aminoethyl ether)-N,N,N′,N′-tetraacetic acid (EGTA) was added in 500mL of this buffer
while 2mM calcium chloride (CaCl2) was added into another 500mL of this buffer. Both
solutions were pre-warmed in a water bath set at 39-40℃ with carbogen.
Fifteen-week-old female C57BL/6J mice were anaesthetised using a cocktail of
ketamine (125mg/kg) and xylazine (25mg/kg) made up in saline. For perfusion, a cannula was
inserted into the inferior vena cava to initiate liver perfusion in situ. The portal vein was
immediately cut to allow excess buffer solution to drain out of the liver. Perfusion was
maintained at 5mL per minute for 15 minutes, after which the liver was digested with
collagenase H (50mL of 1mg/mL; Roche, cat.no. 11074032001) in warm HBSS solution
supplemented with 2mM CaCl2 for 9 minutes. Once perfusion was complete, the liver was
excised, and the digested liver capsule was removed using fine forceps on a glass petri dish
containing HBSS solution. The hepatocytes were then strained through a 100µM cell strainer
and centrifuged 3 times in ice cold CaCl2-HBSS buffer at 50g for 3 minutes at 4℃. Under aseptic conditions, the final cell pellet was resuspended in warm Medium 199 (Sigma-Aldrich,
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Chapter 3 Quantifying the NAD metabolome cat.no. M4530) before seeding onto 6-well plates which were pre-coated with Type 1 rat collagen from tail (Gibco Life Technologies, cat.no. A1048301).
Cells were incubated for 4 hours in a 37℃, 5% CO2 incubator in the following adherence medium: Medium 199 (Sigma-Aldrich, cat.no. M4530) supplemented with penicillin-streptomycin solution (1X p/s) (10,000U/mL, Gibco cat.no. 15140), bovine serum albumin (BSA) (10%, Gibco, cat.no. 15260), UltroSer G (Pall Biosepra, cat.no. 15950-017), dexamethasone (10mM in ethanol, 100nM final concentration, Sigma cat.no. D-4902), and insulin (690µM, 100nM final concentration, Actrapid®). After 4 hours, the adherence media was changed to the following basal media: Medium 199 (Sigma-Aldrich, cat.no. M4530) supplemented with 1% p/s (10,000U/mL, Gibco cat.no. 15140) and dexamethasone (10mM in ethanol, 100nM final concentration, Sigma, cat.no. D-4902). After an overnight incubation in basal media, it was supplemented with the following treatment groups and incubated for 24 hours in a 37℃, 5% CO2 incubator: 1) Control (water), 2) NMN1 (0.2mM), and 3) NMN2
(0.2mM). NMN1 represents labelled NMN with a mass shift of M+6 (labelled at ribose and base of the nicotinyl ring from the nicotinamide moiety) while NMN2 represents labelled NMN with a mass shift of M+7 (labelled at ribose, base of the nicotinyl ring and amide group of the nicotinamide moiety) (Section 2.1.1) Cells were harvested after 24 hours for metabolite extraction (Section 2.2.2), and processing (Section 2.2.3) before undergoing analysis by LC-
MS/MS (Section 2.3) and data analysis (Section 2.4).
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3.2.4 C2C12 and HEK293 Cell Culture
C2C12 and HEK293 cells were grown and maintained in DMEM (Dulbecco's Modified Eagle
Medium) supplemented with 10% v/v fetal bovine serum (FBS) (Life Technologies Australia,
cat.no. 10099141) and 1% v/v penicillin-streptomycin (p/s) solution (Sigma Aldrich, cat.no.
P0781). Cells were seeded at a density of 0.1x106 cells per well (2mL/well) in 6-well plates and incubated for 24 hours in a 37℃, 5% CO2 incubator. Media was then removed and washed
once with warmed 1X phosphate buffered saline (PBS) solution before adding fresh DMEM
media supplemented with either vehicle (water) control or 0.1mM labelled NMN1 (Section
2.1.1). Cells treated at time zero were supplemented with an equivalent amount of unlabelled
NMN as the amount of M+6 labelled NMN1 isotope was limited. Cells were harvested at time points 0, 1, 2, 4, 8, and 24 hours in duplicate and NAD+ metabolites were extracted (Section
2.2.2), and processed (Section 2.2.3) before undergoing analysis by LC-MS/MS (Section 2.3)
and data analysis (Section 2.4). Data is represented as ion intensities normalised to the internal
standard (thymine-d4) and normalised again to the protein concentration of each sample and
represented either as normalised intensities or picomoles per mg protein (pmol/mg protein).
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3.3 Results To optimise detection by tandem mass spectrometry, the MRM transition (mass to charge [m/z] ratios for the precursor [Q1] and product [Q3] ion) for each metabolite,and their respective isotopologues need to be determined. The precursor ion m/z ratio was based on the monoisotopic mass of the unlabelled metabolite, while the fragment/product ion m/z ratio was determined by performing an automatic compound optimisation of mass spectrometry (MS) parameters—declustering potential (DP), collision energy (CE) and cell exit potential (CXP)— to select the m/z ratio with the highest signal intensity. The MRM transition was verified using an online MS/MS spectrum database (pubchem.ncbi.nlm.nih.gov).
Optimising MS parameters is an important process in maximising signal to noise ratios, to obtain well-resolved peaks for each metabolite. The de-clustering potential is the voltage applied to prevent ions in a sample from clustering, which assists the separation of ions of interest in the first quadrupole (Q1) of a triple quadrupole mass spectrometer. The collision energy is the voltage applied to increase the fragment ion intensity during collision induced dissociation (CID) in the second quadrupole, where smaller product ions are generated. The cell exit potential is the voltage applied to assist the transition of the fragmented product ions from the second quadrupole to the third quadrupole (Q3), where the metabolites of interest can be filtered and detected to generate chromatogram peaks.
The greatest advantage of MRM is being able to detect both intact and cleaved versions of the labelled nicotinamide and ribose moieties from NMN. The incorporation of intact labelled nicotinamide and ribose moieties into NAD+ can be measured by detecting a mass shift of either M+6 (from NMN1) or M+7 (from NMN2), while detection of M+1 (labelled nicotinamide from NMN1), M+2 (labelled nicotinamide from NMN2) or M+5 (labelled ribose from either NMN1 or NMN2) indicates incorporation of the cleaved labelled moieties after the glycosidic bond between the nicotinamide and ribose moieties of NMN is broken (Figure 3.2).
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Further, the ability to detect the incorporation of these labelled entities in other NAD+ metabolites, either completely or after the glycosidic bond is broken, allows their contribution to NAD+ biosynthesis to be explored in the context of NMN supplementation.
Figure 3.2 Detecting labelled NMN and NAD+ isotopologues using MRM (multiple reaction monitoring). NAD+ (M+6) and NAD+ (M+7) indicates “complete” incorporation of labelled moieties from M+6 labelled NMN1 and M+7 labelled NMN2, respectively. NAD+ (M+1), NAD+ (M+2) and NAD+ (M+5) indicate “partial” incorporation of labelled moieties after the glycosidic bond between the ribose and nicotinamide is broken. The MRM transitions for each NMN and NAD+ isotopologue are shown in brackets. Red asterisks denote isotope labels and blue dotted lines denote the point of fragmentation. The m/z ratio of the fragment ion detected is shown.
While this is not the first LC-MS/MS method to use MRM to investigate NMN metabolism, it is important to optimise these conditions between LC-MS instruments, buffers and protocols as these MS parameters can vary significantly under various laboratory conditions. Further, the labelled NMN isotopes used in this study were designed specifically to
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trace the metabolic fate of NMN and investigate its influence on the NAD+ metabolome. As
such, the differences in isotope design and application between studies requires optimisation
of MS parameters relevant to the specific metabolites of interest.
3.3.1 Compound optimisation of mass spectrometry parameters for internal standards.
An important component of quantifying analytes by mass spectrometry is the use of internal
standards, which provide a reference point for concentrations. Internal standards are either
structural analogues or stable isotopically labelled (SIL) analogues of the compound of interest.
Suitable candidates are not always available or easily accessible and SIL analogues can often
be expensive. As such, the internal standards, camphorsulfonic acid (CSA), methionine
sulfonate (MES) and thymine-d4 (deuterated), used here, were chosen due to availability, ease
of detection, and because they are not endogenously present in biological samples. Table 3.2
provides details of the optimised compound parameters for these three internal standards.
Table 3.2 MRM transitions for internal standards and mass spectrometry parameters. Internal standard Q1 (m/z) Q3 (m/z) DP (V) CE (V) CXP (V) RT (mins) CSA 230.92 80.00 -170.00 -40.00 -13.00 1.86 MES 196.12 100.00 140.00 31.00 25.00 4.00 Thymine-d4 129.03 42.10 -115.00 -52.00 -11.00 2.00 MRM: multiple reaction monitoring Q1: parent ion, Q3: fragment ion, m/z: mass to charge ratio DP: declustering potential, CE: collision energy, CXP: collision cell exit potential V: volts, RT: retention time CSA: camphorsulfonic acid, MES: methionine sulfonate
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3.3.2 Optimisation of mass spectrometry parameters for NAD+ metabolites
Following the internal standards, the optimisation of MRM parameters was repeated for
analytes of the NAD+ metabolome that were relevant to this investigation. The optimised
compound parameters for unlabelled (M+0) NAD+ metabolites are detailed in Table 3.3 for the following eight compounds - NAM, NMN, NR, NAD+, NA, NaR, NaAD and NaMN. Note
that isotopologues (isotope labelled versions) of a given metabolite have different Q1 and Q3
depending on the labelled state (i.e., mass shift), but share the same depolarisation potential
(DP), collision energy (CE), cell exit potential (CXP) and retention time (RT). The MRM
transitions and their mass spectrometry parameters are detailed in Table 3.4 for amidated
metabolites and in Table 3.5 for deamidated metabolites.
Table 3.3 MRM transitions and MS parameters for unlabelled NAD+ metabolites Isotopologue Q1 (m/z) Q3 (m/z) DP CE CXP RT (precursor_product) ion (V) (V) (V) (mins) mass shift NAM_0_0 122.80 79.80 80.00 30.00 25.00 1.99 NMN_0_0 335.15 123.05 48.00 24.00 11.00 22.50 NR_0_0 255.00 123.03 64.00 30.00 13.00 12.50 NAD+_0_0 664.00 428.09 33.00 36.00 31.00 16.70 NA_0_0 123.80 77.80 70.00 25.00 10.00 4.30 NaR_0_0 256.20 124.20 41.00 27.00 6.00 10.50 NaAD_0_0 665.00 428.10 139.00 35.00 38.00 17.05 NaMN_0_0 336.10 123.80 66.00 28.00 11.00 20.90 Abbreviations: NAM: nicotinamide, NMN: nicotinamide mononucleotide, NR: nicotinamide riboside and NAD+: nicotinamide adenine dinucleotide (oxidised), NA: nicotinic acid, NaR: nicotinic acid riboside, NaAD: nicotinic acid adenine dinucleotide, NaMN: nicotinic acid mononucleotide
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Table 3.4 MRM transitions and mass spectrometry parameters for isotopologues of NAM, NMN, NR and NAD+ Isotopologue Q1 (m/z) Q3 (m/z) DP (V) CE (V) CXP (V) RT (mins) (precursor_product) ion mass shift NAM_0_0 122.80 79.80 80.00 30.00 25.00 1.99 NAM_1_0 123.80 79.80 80.00 30.00 25.00 1.99 NAM_1_1 123.80 80.80 80.00 30.00 25.00 1.99 NAM_2_1 124.80 80.80 80.00 30.00 25.00 1.99 NMN_0_0 335.15 123.05 48.00 24.00 11.00 22.50 NMN_1_1 336.15 124.05 48.00 24.00 11.00 22.50 NMN_2_2 337.15 125.05 48.00 24.00 11.00 22.50 NMN_5_0 340.15 123.05 48.00 24.00 11.00 22.50 NMN_6_1 341.15 124.05 48.00 24.00 11.00 22.50 NMN_7_2 342.15 125.05 48.00 24.00 11.00 22.50 NR_0_0 255.00 123.03 64.00 30.00 13.00 12.50 NR_1_1 256.00 124.03 64.00 30.00 13.00 12.50 NR_2_2 257.00 125.03 64.00 30.00 13.00 12.50 NR_5_0 260.00 123.03 64.00 30.00 13.00 12.50 NR_6_1 261.00 124.03 64.00 30.00 13.00 12.50 NR_7_2 262.00 125.03 64.00 30.00 13.00 12.50 NAD+_0_0 664.00 428.09 33.00 36.00 31.00 16.70 NAD+_1_0 665.00 428.09 33.00 36.00 31.00 16.70 NAD+_2_0 666.00 428.09 33.00 36.00 31.00 16.70 NAD+_5_0 669.00 428.09 33.00 36.00 31.00 16.70 NAD+_6_0 670.00 428.09 33.00 36.00 31.00 16.70 NAD+_7_0 671.00 428.09 33.00 36.00 31.00 16.70 NAD+_1_1 665.00 429.09 33.00 36.00 31.00 16.70 NAD+_2_2 666.00 430.09 33.00 36.00 31.00 16.70 NAD+_5_5 669.00 433.09 33.00 36.00 31.00 16.70 NAD+_6_1 670.00 429.09 33.00 36.00 31.00 16.70 NAD+_6_6 670.00 434.09 33.00 36.00 31.00 16.70 NAD+_7_0 671.00 428.09 33.00 36.00 31.00 16.70 NAD+_7_1 671.00 429.09 33.00 36.00 31.00 16.70 NAD+_7_7 671.00 435.09 33.00 36.00 31.00 16.70 NAD+_8_1 672.00 429.09 33.00 36.00 31.00 16.70 NAD+_10_5 674.00 433.09 33.00 36.00 31.00 16.70 NAD+_11_5 675.00 433.09 33.00 36.00 31.00 16.70 NAD+_11_6 675.00 434.09 33.00 36.00 31.00 16.70 NAD+_12_5 676.00 433.09 33.00 36.00 31.00 16.70 NAD+_12_6 676.00 434.09 33.00 36.00 31.00 16.70 NAD+_13_6 677.00 434.09 33.00 36.00 31.00 16.70 Abbreviations: NAM: nicotinamide, NMN: nicotinamide mononucleotide, NR: nicotinamide riboside and NAD+: nicotinamide adenine dinucleotide (oxidised).
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Table 3.5 MRM transitions and mass spectrometry parameters for isotopologues of NA, NaR, NaMN and NaAD Isotopologue Q1 (m/z) Q3 (m/z) DP (V) CE (V) CXP (V) RT (mins) (precursor_product) ion mass shift NA_0_0 123.80 77.80 70.00 25.00 10.00 4.30 NA_1_1 124.80 78.80 70.00 25.00 10.00 4.30 NaR_0_0 256.20 124.20 41.00 27.00 6.00 10.50 NaR_1_1 257.20 125.20 41.00 27.00 6.00 10.50 NaR_5_0 261.20 124.20 41.00 27.00 6.00 10.50 NaR_6_1 262.20 125.20 41.00 27.00 6.00 10.50 NaAD_0_0 665.00 428.10 139.00 35.00 38.00 17.05 NaAD_1_0 666.00 428.10 139.00 35.00 38.00 17.05 NaAD_5_0 670.00 428.10 139.00 35.00 38.00 17.05 NaAD_6_0 671.00 428.10 139.00 35.00 38.00 17.05 NaAD_1_1 666.00 429.10 139.00 35.00 38.00 17.05 NaAD_2_1 667.00 429.10 139.00 35.00 38.00 17.05 NaAD_5_5 670.00 433.10 139.00 35.00 38.00 17.05 NaAD_6_1 671.00 429.10 139.00 35.00 38.00 17.05 NaAD_6_5 671.00 433.10 139.00 35.00 38.00 17.05 NaAD_6_6 671.00 434.10 139.00 35.00 38.00 17.05 NaAD_7_1 672.00 429.10 139.00 35.00 38.00 17.05 NaAD_7_6 672.00 434.10 139.00 35.00 38.00 17.05 NaAD_7_7 672.00 435.10 139.00 35.00 38.00 17.05 NaAD_10_5 675.00 433.10 139.00 35.00 38.00 17.05 NaAD_11_5 676.00 433.10 139.00 35.00 38.00 17.05 NaAD_11_6 676.00 434.10 139.00 35.00 38.00 17.05 NaAD_12_6 677.00 434.10 139.00 35.00 38.00 17.05 NaMN_0_0 336.10 123.80 66.00 28.00 11.00 20.90 NaMN_1_1 337.10 124.80 66.00 28.00 11.00 20.90 NaMN_5_0 341.10 123.80 66.00 28.00 11.00 20.90 NaMN_6_1 342.10 124.80 66.00 28.00 11.00 20.90 Abbreviations: NA: nicotinic acid, NaR: nicotinic acid riboside, NaAD: nicotinic acid adenine dinucleotide, NaMN: nicotinic acid mononucleotide.
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The LC-MS method was not optimised for recovery however, to compensate for
potential losses during sample handling and processing, samples and standards were subjected
to the same extraction method. R-squared regression (R2) values for each metabolite are shown
in Table 3.6 for accuracy, along with Relative Standard Deviation (RSD) for measures of precision. The Limit of Quantification (LOQ), the lowest concentration at which the metabolite can be reliably detected, is a measure of sensitivity of the LC-MS method. LOQ is calculated from the standard deviation of the calibration curve generated for each metabolite and also shown in Table 3.6. The LOQ and RSD for NAD+ metabolites, especially NR, is relatively high compared to previous studies (Evans et al. 2010; Trammell and Brenner 2013) however,
NMN and NR are known to be unstable metabolites in mouse serum and blood (Liu et al. 2018) contributing to the high variability detected in our samples in vivo (Chapter 5).
Table 3.6 LC-MS precision and accuracy parameters for each metabolite Metabolite Transition CE RT LOQ R2 RSD (m/z) (V) (mins) (pmol) (%) NMN 335.15>123.05 24.00 18.80 91.01 0.998 8.10 NaMN 336.10>123.80 28.00 18.50 92.92 0.997 10.10 NR 255.00>123.03 13.00 11.45 44529.13 0.989 24.80 NaR 256.20>124.20 6.00 10.20 180.77 0.999 7.00 NaAD 665.00>428.10 35.00 16.80 85.82 0.985 20.90 Nam 122.80>79.80 30.00 1.98 87.91 0.991 17.10 Na 123.80>77.80 25.00 4.20 91.50 0.999 5.80 NAD+ 664.00>428.09 36.00 16.95 89.35 0.989 11.50 Abbreviations: CE (V): collision energy (volts); RT (mins): retention time (minutes); LOQ (pmol): level of quantification (picomoles); R2: R-squared regression; RSD (%): relative standard deviation; NA: nicotinic acid, NaAD: nicotinic acid adenine dinucleotide, NAD+: nicotinamide adenine dinucleotide (oxidised), NAM: nicotinamide, NaMN: nicotinic acid mononucleotide. NaR: nicotinic acid riboside, NMN: nicotinamide mononucleotide, and NR: nicotinamide riboside.
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3.3.3 Detecting labelled NMN isotopes using LC-MS/MS and MRM.
Following the custom synthesis of strategically designed NMN isotopes (see Chapter 2, Section
2.1.1), the purity and mass of these compounds were verified to ensure proper labelling using
the optimised LC-MS/MS method and their predetermined MRM transitions. Representative
chromatograms are shown for unlabelled NMN (Figure 3.3 A) and the two labelled NMN isotopes, NMN1 (M+6) (Figure 3.3 B) and NMN2 (M+7) (Figure 3.3 C). These compounds appeared at their expected m/z, with the appearance of smaller peaks from other isotopologues that can be attributed to the natural abundance of 13C (1.1%) and 15N (0.37%) isotope labels
(Deléens, Cliquet, and Prioul 1994; Dijkstra et al. 2006; Yu et al. 2019) as well as some
impurity or degradation of the NMN isotopes over time, especially NMN2 (M+7) where there
is a significant M+6 peak visible (in yellow) which represents the substitution of amide-15N with amide-14N from the nicotinamide moiety.
Figure 3.3 Representative chromatograms of unlabelled (M+0) and labelled NMN isotopes from injection of a 100 µM standard solution. (A) Unlabelled NMN (M+0) with MRM transition 335.15>123.05. (B) NMN (M+6) or NMN1 with MRM transition 341.15>124.05. (C) NMN (M+7) or NMN2 with MRM transition 342.15>125.05.
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3.3.4 Tracing the metabolic fate of NMN isotopes using LC-MS/MS to detect labelled
NAD+ metabolites.
To test this LC-MS/MS method in a biologically relevant sample, primary hepatocytes were
isolated from fifteen-week-old female C57BL/6 mice and treated with the labelled NMN
isotopes NMN1 and NMN2 for 24 hours. As the liver contains one of the highest concentrations
of NAD+ and is highly active in the metabolism of orally delivered NAD+ precursors (Liu et
al. 2018), primary hepatocytes serve as a good in vitro model to explore NAD+ metabolism
following supplementation with NMN. The labelled nicotinamide (15N) and ribose (13C)
moieties were traced in NAD+ metabolites involved in the canonical pathway for NMN
metabolism, the salvage pathway.
NMN synthesises NAD+ via the three isoforms of nicotinamide mononucleotide
adenylyltransferase (NMNAT1-3) (Raffaelli et al. 2002; Schweiger et al. 2001; Zhang et al.
2003) before it is degraded by NAD+-consuming enzymes such as the sirtuins, poly-ADP-
ribose polymerases (PARPs) and glycohydrolases (CD38/CD157) into nicotinamide (NAM)
and ADP-ribose (ADPr) (Imai et al. 2000a; Kim, Jacobson, and Jacobson 1993; Kim, Zhang, and Kraus 2005).
ADPr is an important by-product of NAD+ consumption, where protein deacetylation
by sirtuin enzymes results in the transfer of the acetyl group from acetylated protein substrates
to form O-acetyl-ADP-ribose (OAADPr) (Tanny and Moazed 2001), acting as a signalling
molecule for various proteins such as the Ca2+ channel TRPM2 (Tanner et al. 2000; Tong and
Denu 2010; Zhao et al. 2004). Generation of ADPr is also important for PARPs in protein and
histone ADP-ribosylation during DNA damage repair (Palazzo et al. 2018) and by
glycohydrolases such as CD38/CD157 to regulate intracellular calcium signalling (Deshpande
et al. 2003; Howard et al. 1993) and the immune response (Partida-Sánchez et al. 2001). NAM
is on the other hand recycled back into the salvage pathway to synthesise NMN via
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nicotinamide phosphoribosyltransferase (NAMPT), the rate limiting enzyme in this pathway
(Revollo, Grimm, and Imai 2004; Wang et al. 2006b), which is then used to synthesise NAD+
via NMNAT1-3 enzymes (Berger et al. 2005; Garavaglia et al. 2002; Schweiger et al. 2001;
Zhang et al. 2003).
As expected, intact labelled NMN1 with an overall mass shift of M+6 (Figure 3.4 A)
and intact labelled NMN2 with an overall mass shift of M+7 (Figure 3.4 B) was detected in
cells treated with NMN1 and NMN2, respectively. To avoid any residual NMN1 or NMN2
supplemented in the cell culture media affecting the measurement of intracellular metabolites,
cells were rinsed with phosphate buffered saline (1XPBS) and discarded prior to cell harvest
in extraction buffer. As a direct precursor to NAD+ biosynthesis, supplementation with labelled
NMN compounds led to increased levels of intracellular NAD+ which incorporated the M+6
and M+7 mass shifts from NMN1 and NMN2, respectively (Figure 3.4 C-D). The presence of these labels in NAD+ indicates these metabolites incorporated the ribose and nicotinamide
moieties intact, as both moieties were labelled with 13C and 15N isotopes to contribute to the
overall increase in mass shift.
As expected, we also detected labelling of the NAD+ degradation product, nicotinamide
(NAM), as NAM (M+1) (Figure 3.5 A) and NAM (M+2) (Figure 3.5 B) following
supplementation with NMN1 and NMN2 respectively. These represent the cleavage activity of
NAD+ (M+6) and NAD+ (M+7), respectively, by NAD+-consuming enzymes. These data do not exclude the possibility that NMN undergoes intracellular cleavage at the glycosidic bond to release free NAM and ribose-5-phosphate, as this bond could be labile under biological conditions (Bell, Yeates, and Eisenberg 1997; Göckel and Richert 2015; Liu et al. 2008).
Following cleavage of NAD+, the ADP-ribose moiety is liberated through NAD+-consuming
activities while NAM is recycled back into NMN.
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The regeneration of NMN from NAM via the NAD+ salvage pathway involving the rate-limiting enzyme, NAMPT (Revollo, Grimm, and Imai 2004), was detected. This was inferred based on the presence of NMN (M+1) (Figure 3.5 C) and NMN (M+2) (Figure 3.5
D) which form the precursors for the synthesis of NAD+ (M+1) (Figure 3.5 E) and NAD+
(M+2) (Figure 3.5 F), respectively, via NMNAT1-3 (Raffaelli et al. 2002; Schweiger et al.
2001; Zhang et al. 2003).
These results largely verified that the labelling strategy using NMN isotopes to trace labelled nicotinamide and ribose moieties in NAD+ metabolites through the optimised LC-
MS/MS method developed here, are sound. These NMN isotopes demonstrated expected behaviour through the canonical salvage pathway in a biologically relevant in vitro model, ensuring the use of these strategies in subsequent experiments to explore NMN metabolism.
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Figure 3.4 Labelled NMN compounds are detected intact and contribute to an increase in NAD+ in primary hepatocytes. Primary hepatocytes were isolated from 15-week-old female C57BL/6J mice and treated with either 1) vehicle control, 2) NMN1 (0.2 mM), or 3) NMN2 (0.2 mM). Data are represented as mean ± s.d. (n=3) and normalised to the average of internal standards (MES, CSA and thymine-d4). (A) Increase in NMN (M+6) detected in cells after extracellular NMN1 supplementation. (B) Increase in NMN (M+7) detected in cells after extracellular NMN2 supplementation. (C) Increase in NAD+ (M+6) detected in cells after extracellular NMN1 supplementation. (D) Increase in NAD+ (M+7) detected in cells after extracellular NMN2 supplementation.
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Figure 3.5 Labelled NAD+ is degraded and contributes to increases in metabolites in the NAD+ recycling pathway. Primary hepatocytes were isolated from 15-week-old female C57BL/6J mice and treated with either 1) vehicle control, 2) NMN1 (0.2 mM), or 3) NMN2 (0.2 mM). Data are represented as mean ± s.d. (n=3) and normalised to the average of internal standards (MES, CSA and thymine-d4). (A) NAM (M+1) formed from cleavage of NAD+ (M+6) by NAD+-consuming enzymes. (B) NAM (M+2) formed from cleavage of NAD+ (M+7) by NAD+-consuming enzymes. (C) NMN (M+1) synthesised from NAM (M+1) via enzyme, NAMPT. (D) NMN (M+2) synthesised from NAM (M+2) via enzyme NAMPT. (E) NAD+ (M+1) synthesised from NMN (M+1) via enzyme isoforms NMNAT1-3. (F) NAD+ (M+2) synthesised from NMN (M+2) via enzyme isoforms NMNAT1-3.
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3.3.5 Intracellular NR is detected following extracellular supplementation of NMN in
primary hepatocytes.
The mechanism for transport of NMN into cells remains elusive. While there is evidence that
NMN is transported intact (Grozio et al. 2019a; Yoshino et al. 2011), others argue NMN
requires dephosphorylation into nicotinamide riboside (NR) for transport and utilisation inside
the cell (Ratajczak et al. 2016). To investigate mechanisms of NMN transport into cells, the
incorporation of the labelled nicotinamide (15N) and ribose (13C) moieties into NMN and NR isotopologues were traced in primary hepatocytes.
Initially, the intact labelling of NMN (M+6) (Figure 3.4 A) and NMN (M+7) (Figure
3.4 B) detected in primary hepatocytes after 24 hours exposure to NMN1 and NMN2, respectively, suggested these isotopes were transported directly into the cell. The concomitant detection of NR (M+6) (Figure 3.6 A) and NR (M+7) (Figure 3.6 B), also suggested that these
NMN isotopes underwent dephosphorylation into NR and were taken up by cells. Direct transport of NMN into cells via SLC12A8 has been reported to involve rapid uptake (within 1 minute) and immediate conversion (within 15 minutes) to NAD+ (Grozio et al. 2019a; Mills et
al. 2016). Compared to this minute-order uptake of NMN into the cell reported by Grozio and
colleagues, the extracellular dephosphorylation of NMN to NR followed by its intracellular re-
phosphorylation back into NMN reportedly occurs slowly over the course of 24 hours (Belenky
et al. 2007; Ratajczak et al. 2016) which is consistent with the detection of NMN at this time
point observed in this study.
In order to compare the intracellular labelled pools of NMN and NR, the fold change
increases in labelled NMN and NR species were calculated relative to the control group.
Following exposure to NMN1, the fold change increase in M+6 labelled NR relative to control
was 968.84, while the fold change increase in M+6 NMN relative to control was 639.22
(Figure 3.6 C). These data indicate NR is more abundant than NMN in the cell following
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supplementation with NMN1, which suggests that NMN transport may occur via
dephosphorylation to NR. Similarly, NMN2 supplementation led to a fold change increase in
M+7 NR of 1276.92 relative to control (Figure 3.6 D). Despite this being lower than the fold
change increase in M+7 NMN of 2503.17 relative to control, this still suggests some evidence
that NR is being produced abundantly following NMN2 supplementation. These data alone are
not enough evidence to prove this mechanism, nor does this exclude the possibility that NMN
may still achieve cellular entry through direct intact transport. This is especially considering
the fact that intracellular labelled NMN was observed following administration with labelled
NMN isotopes in this study, an observation which contradicts findings from a similar study by
Ratajczak and colleagues where labelled NMN was not detected in primary hepatocytes
supplemented with labelled NMN (2H-13C-NMN) over a 6-hour time course. Considering the longer time point (24 hours) captured in this study, the detection of labelled NMN could come from its conversion to NR inside the cell, which is a possibility supported by the abundant detection of labelled NR observed in this study. This is investigated more closely later in this chapter, in the mammalian cell lines C2C12 and HEK293, where labelled NMN is supplemented in the cell culture media and harvested at multiple time points.
Overall, without completely excluding the potential uptake of NMN directly through the NMN transporter, the abundance of labelled NR detected in primary hepatocytes following exogenous supplementation with labelled NMN isotopes suggest that NMN may be taken up through dephosphorylation into NR prior to its uptake and subsequent re-phosphorylation to
NMN once inside the cell.
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Figure 3.6 An abundance of intact labelled NR is detected following supplementation with labelled NMN. Primary hepatocytes were isolated from 15-week-old female C57BL/6J mice and treated with either 1) vehicle control, 2) NMN1 (0.2 mM), or 3) NMN2 (0.2 mM). Data are represented as mean ± s.d. (n=3) and normalised to the average of internal standards (MES, CSA and thymine-d4). (A) Intact labelled NR (M+6) following supplementation with labelled NMN (0.2 mM). (B) Intact labelled NR (M+7) following supplementation with labelled NMN (0.2 mM). (C) Fold change of M+6 NMN and NR relative to control following supplementation with labelled NMN1 or NMN2 (0.2 mM). (D) Fold change of M+7 NMN and NR relative to control following supplementation with labelled NMN1 or NMN2 (0.2 mM).
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3.3.6 NMN does not lead to increases in deamidated metabolites, NaMN and NaAD,
following supplementation in primary hepatocytes.
These findings thus far, demonstrate that NMN metabolism occurs through the expected
salvage pathway, primarily involving amidated NAD+ metabolites. That is, NAD+ is
synthesised inside the cell and subsequently degraded into nicotinamide (NAM) where it is
recycled back into the salvage pathway to synthesise NMN via NAMPT, and NAD+, via
NMNAT1-3. The effect of extracellular NMN on intracellular NR was also demonstrated in
primary hepatocytes through a higher abundance of labelled NR pool versus labelled NMN
detected inside the cell, suggesting the potential mechanism for the uptake of NMN may
involve dephosphorylation to NR.
It is unknown whether exogenous NMN treatment influences the deamidated NAD+
metabolite nicotinic acid mononucleotide (NaMN), nicotinic acid adenine dinucleotide
(NaAD), nicotinic acid (NA) and nicotinic acid riboside (NaR) as there are no established
mammalian pathways for this to be expected, let alone investigated. These deamidated
metabolites form another NAD+ biosynthesis pathway called the Preiss-Handler pathway.
NaMN is synthesised either from NA, via nicotinic acid phosphoribosyltransferase (NaPRT), or from NaR, via nicotinamide riboside kinases (NRK1/2), the same enzymes responsible for the conversion of NR to NMN in the salvage pathway (Bieganowski and Brenner 2004; Tempel et al. 2007). NaMN is then converted into NaAD via NMNAT1-3, which also convert NMN to
NAD+ in the salvage pathway. The glutamine-dependent reaction catalysed by NAD+
synthetase (NADSYN1) is then used to amidate NaAD to NAD+ in the final step of the Preiss-
Handler pathway.
It is important to recognise that none of the precursors from the salvage pathway, such
as NMN or NR, are involved in the biosynthesis of NAD+ in this pathway. Despite this, the
presence of the deamidated metabolite, NaAD, was strikingly detected following
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supplementation with the amidated metabolite, nicotinamide riboside (NR) (Trammell et al.
2016a), suggesting an unknown mammalian pathway may exist that forms NaAD from
metabolites in the salvage pathway such as NMN and NR.
To explore this possibility, the abundance of these deamidated metabolites were
explored following supplementation with NMN isotopes, focusing on NaMN and NaAD.
Despite our best efforts (details described in following paragraphs), our results showed that
levels of NaMN and NaAD was not increased, nor labelled, following exogenous
supplementation with labelled NMN. From these findings, it appears that the deamidation
pathway does not occur in primary hepatocytes.
Interpretation of mass spectrometry data obtained from NaAM and NaAD presented
some challenges due to the isobaric (differ by one Dalton) nature to their amidated counterparts,
NMN and NAD+, respectively. The monoisotopic mass of NaMN (335.04 g/mol; PubChem
CID 5288991) and NaAD (665.10 g/mol; PubChem CID 165491) is one Dalton higher than
that of NMN (334.06 g/mol; PubChem CID 14180) and NAD+ (664.11 g/mol; PubChem CID
5893), respectively. Consequently, the isotope labelling strategy, where an extra isotope label
is incorporated, will cause several pairs of NaMN/NMN and NaAD/NAD+ isotopologues to
share the same MRM transitions. For example, the MRM transitions for NMN labelled at the
nicotinyl ring base of the nicotinamide moiety, NMN (M+1), with m/z 336.15>124.05, will
have the same MRM transition as unlabelled NaMN (M+0), with m/z 336.10>123.80.
Similarly, the isotopologue pairs NMN (M+2)/NaMN (M+1), NMN (M+6)/NaMN (M+5) and
NMN (M+7)/NaMN (M+6) will also share the same MRM transitions (Figure 3.7) causing
them to be detected under the same chromatogram peak or in some cases overlap. This can be
observed in Figure 3.3 B and C where the smaller peaks to the left of the main NMN peaks represent the corresponding isotopologues from NaMN.
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To determine which isotopologue each peak truly represents, the chromatogram peaks
from primary hepatocytes supplemented with unlabelled NMN were explored. Figure 3.7 A
shows chromatogram peaks following supplementation with unlabelled NMN (0.2 mM) while
Figure 3.7 B shows chromatogram peaks following supplementation with M+6 labelled
NMN1 (0.2 mM) in primary hepatocytes. As shown in Figure 3.7 A, the larger main peak (in
dark blue) represents NMN (M+0) while the smaller peak (in light blue) represents both NMN
(M+1) and NaMN (M+0) under the same peak. As these cells were supplemented with
unlabelled NMN, the NMN (M+1) detected here represents the incorporation of isotope labels
that exist normally in nature, that is, the naturally occurring isotope 13C occurs ~1.1% and 15N occurs ~0.37% in natural abundance (Deléens, Cliquet, and Prioul 1994; Dijkstra et al. 2006;
Yu et al. 2019). The isotope distributions for NMN (C11H15N2O8P) can be calculated with the
isotope distribution calculator available at https://www.sisweb.com/mstools/isotope.htm. The natural abundance of NMN with an extra isotope (MW 335) is 13.2% (Figure 3.7 C), which is approximately one tenth (or 0.1) of the abundance of unlabelled NMN (MW 334). Any abundance detected above this ratio, therefore, represents the formation of unlabelled NaMN
(M+0) following supplementation with unlabelled NMN which was quantified in Figure 3.7
D. The ratio of NMN (M+1) (mean 37.26) to NMN (M+0) (mean 413.78) was 0.09 and the ratio of NaMN (M+0) (mean 25.20) to NMN (M+0) was 0.06. As both were less than 0.1, it was established that the combined NMN (M+1)/NaMN (M+0) peak most likely represented
NMN (M+1). Further, very little unlabelled NaMN was endogenously present or newly synthesised following supplementation with unlabelled NaMN. As such, this negligible influence of NMN on NaMN levels indicated that the shared peaks between pairs of NMN and
NaMN isotopologues in Figure 3.7 B most likely represent the NMN isotopologues rather than their NaMN counterparts.
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Further, following supplementation with the NMN isotope NMN1, the abundance of the NMN (M+7)/NaMN (M+6) peak was similar to the control (no NMN), indicating that not only was there very little NMN (M+7) detected, very little NaMN (M+6) was present in samples. Therefore, the abundance of labelled NaMN (M+6) observed after supplementing with the labelled NMN isotope NMN2 (M+7) most likely represents the abundance of NMN
(M+7), rather than NaMN (M+6), which shares a similar MRM transition.
These results show that levels of the deamidated metabolite NaMN are largely not affected by exogenous supplementation with NMN.
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Figure 3.7 Chromatogram peaks of NMN/NaMN isotopologue pairs sharing the same MRM transition. Primary hepatocytes were isolated from 15-week-old female C57BL/6J mice and treated with either unlabelled NMN (0.2 mM) or NMN1 (0.2 mM). Chromatogram peaks of unlabelled and labelled NMN/NaMN isotopologues after supplementation with (A) unlabelled, (B) M+6 labelled NMN1 and quantified in (D) and (E), respectively. (C) The isotope distribution of NMN (C11H15N2O8P) was calculated with the isotope distribution calculator available at https://www.sisweb.com/mstools/isotope/htm. Data are represented as mean ± s.d. (n=3) and normalised to the average of internal standards (MES, CSA and thymine-d4).
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In contrast to the chromatograms shared by NMN/NaMN pairs of isotopologues
described in Figure 3.7 (above), the normalised intensities calculated from the integration of
chromatogram peaks shared by NAD+/NaAD pairs of isotopologues showed discrepancies. The
natural abundance of M+1 labelled NAD+ is 26.2% (Figure 3.8 C, below) and any value above this percentage in an injected sample suggests the presence of unlabelled NaAD (M+0) as it shares the same MRM transition as M+1 labelled NAD+ (665.00>428.09 for NAD+ [M+1] vs.
665.00>428.10 for NaAD [M+0]).
This was the case observed in Figure 3.8 A, where the chromatogram peak shared by
NAD+ (M+1) and NaAD (M+0) was approximately half (~50%) of the unlabelled NAD+ (M+0)
peak (absolute percentage not quantified), suggesting NaAD is detected in cells following
supplementation with NMN. The same observation was made following supplementation with
M+6 labelled NMN1 (Figure 3.8 B), although this value was slightly lower (absolute
percentage not quantified) but still above the expected 26.2% from natural abundance. The
autogenerated values which were extracted following the integration of chromatogram peaks
suggest that NaAD is detected in very low amounts in samples following supplementation with
NMN, and is independent of the increases belonging to corresponding NAD+ isotopologues.
Following supplementation with unlabelled NMN, the combined normalised intensities
calculated from NAD+ (M+1) and NaAD (M+0) is approximately 18.1% of the intensity
calculated from NAD+ (M+0) (Figure 3.8 D), of which NAD+ (M+1) contributes the majority and most likely represents naturally abundant isotope. Following supplementation with M+6 labelled NMN1, M+6 labelled NAD+ increases but NaAD (M+5), which shares the same MRM
transition, does not (Figure 3.8 E). Similarly, NAD+ (M+1) increases slightly whereas NaAD
(M+0) does not, suggesting that NaAD is not present in samples. The isotopologue pairs NAD+
(M+7)/NaAD (M+6) and NAD+ (M+2)/NaAD (M+1) represent complete and partial
incorporation of labelled moieties from M+7 labelled NMN2 respectively, though as expected,
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NMN1. These data suggest that, like NaMN, levels of the deamidated metabolite NaAD is not detected in primary hepatocytes following treatment with exogenously supplemented NMN.
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Figure 3.8 Chromatogram peaks from NAD+/NaAD isotopologues sharing the same MRM transition. Primary hepatocytes were isolated from 15-week-old female C57BL/6J mice and treated with either unlabelled NMN (0.2 mM) or NMN1 (0.2 mM). Chromatogram peaks of unlabelled and labelled NMN/NaMN isotopologues after supplementation with (A) unlabelled or (B) M+6 labelled NMN1 + and quantified in (D) and (E), respectively. (C) The isotope distribution of NAD (C21H27N7O14P2) was calculated with the isotope distribution calculator available at https://www.sisweb.com/mstools/isotope/htm. Data are represented as mean ± s.d. (n=3 technical replicates) and normalised to the average of internal standards (MES, CSA and thymine-d4).
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3.3.7 Time-course tracing of NMN metabolism in C2C12 and HEK293 cells – amidated metabolites
The study conducted in primary hepatocytes achieved two things—first, it verified the labelling
strategy of the labelled NMN isotopes NMN1 and NMN2, and second, it validated the use of
LC-MS/MS in MRM mode optimized to detect unlabeled and labelled NAD+ metabolites in a
biologically relevant in vitro model. However, as we only used a single time point, we are
unable to evaluate the dynamics of NMN assimilation, such as how fast NMN is converted into
NAD+. As such, in this next study, samples were harvested at multiple time points—0, 1, 2, 4,
8, and 24 hours—to profile any potential meaningful incorporation of labelled moieties from
NMN into NAD+ metabolites over time. Based on differences in enrichment rates, we can infer which NAD+ metabolite pools are proximal to exogenous NMN and have faster enrichment.
Further, while primary cells are isolated directly from tissues, and are closer in
morphology and function than cells from cultured cell lines, they have limited proliferative
capacity and are overall more difficult to obtain and maintain. Thus, the mouse myoblast cell
line, C2C12, and the human embryonic kidney cell line, HEK293, were used. C2C12 were
used as muscle is an important site of action for NAD+ precursors (Agerholm et al. 2017; Ryu
et al. 2016), while HEK293 cells represent an easy-to-use non-differentiating cell line for investigating NAD+ metabolism (Cantó et al. 2012; Davila et al. 2018; Kulikova et al. 2019).
The labelled NMN isotope NMN1, was added to culture media at a final concentration of 0.1
mM, and harvested at time points, 1, 2, 4, 8, and 24 hours. For the time zero sample, we used
an equivalent dose of NMN, but unlabeled, to conserve our labelled substrate. NMN2 labelling
experiments were not performed due to limited amount of substrates available.
Intact, labelled NMN1 (M+6) was detected in C2C12 and HEK293 cells within the first
hour as shown in Figure 3.9 A and Figure 3.9 B respectively, indicating rapid uptake into both
cell lines. The levels of NMN detected were either maintained (in C2C12 myoblasts) or
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Chapter 3 Quantifying the NAD metabolome increased (in HEK293 cells) by the 8-hour time point, but showed a steep decline by the 24- hour time point in both cell lines, suggesting that by this point, NMN in the media has been depleted. This was confirmed by observing the labelling pattern of (M+6) labelled NAD+, which increased over time in both cell lines as shown in Figure 3.9 C for C2C12 and Figure
3.9 D for HEK293 cells, suggesting that while transport of NMN into the cell may be rapid
(within 1 hour), conversion into NAD+ tends to occur gradually.
The consumption of NAD+ , which produces nicotinamide, was also observed over time indicated by the detection of labelled NAM (M+1) in C2C12 (Figure 3.10 A) and HEK293
(Figure 3.10 B). ). However, the appearance of NAM (M+1) could be attributed to cleavage of
NMN (M+6) or NR (M+6) through a labile glycosidic bond as NR levels were quite high compared to NAD+ (M+6) at the same time points. Interestingly, the formation of NMN (M+1) from its precursor NAM (M+1), was low in both C2C12 (Figure 3.10 C) and HEK293 (Figure
3.10 D) cells, with minimum changes observed over time which may suggest the NMN pool is rapidly converted to NAD+ by fast acting NMNAT enzymes (Berger et al. 2005; Garavaglia et al. 2002; Schweiger et al. 2001; Zhang et al. 2003), which may explain why very little fluctuation in this intermediate species was observed.
In contrast, synthesis of NAD+ (M+1) (Figure 3.10 E-F) from NMN (M+1) increased over time in both cell lines, indicating activation of the salvage pathway following exogenous supplementation with NMN to maintain endogenous NAD+ levels. Interestingly, M+6 intact labelling of NR was detected in C2C12 (Figure 3.9 E) and HEK293 (Figure 3.9 F) cells, similar to that observed in primary hepatocytes (Figure 3.6 A) following extracellular supplementation of NMN1, providing more evidence to suggest its dephosphorylation into NR via CD73 (Grozio et al. 2013).
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Figure 3.9 Labelled NMN detected intact and contributes to increases in NAD+ and NR over time in cell. C2C12 and HEK293 cells were supplemented with labelled NMN (M+6) (0.1mM) for 1, 2, 4, 8, and 24 hours. An equal volume of unlabelled NMN (0.1mM) was supplemented at time zero and harvested for LC-MS/MS analysis. Data represent picomoles normalised to mg protein (pmol/mg protein) with mean ± s.d. (n=2). (A)-(B) Labelled (M+6) NMN in C2C12 and HEK293 cells. (C)-(D) Labelled (M+6) NAD+ in C2C12 and HEK293 cells. (E)-(F) Labelled (M+6) NR in C2C12 and HEK293 cells.
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Figure 3.10 Degradation of NAD+ and nicotinamide salvage C2C12 and HEK293 cells were supplemented with labelled NMN (M+6) (0.1mM) for 1, 2, 4, 8, and 24 hours. An equal volume of unlabelled NMN (0.1mM) was supplemented at time zero and harvested for LC-MS/MS analysis. Data represent picomoles normalised to mg protein (pmol/mg protein) with mean ± s.d. (n=2). (A)-(B) Labelled (M+1) NAM in C2C12 and HEK293 cells. (C)-(D) Labelled (M+1) NMN in C2C12 and HEK293 cells. (E)-(F) Labelled (M+1) NAD+ in C2C12 and HEK293 cells.
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3.3.8 Time-course tracing of NMN metabolism in C2C12 and HEK293 cells – deamidated
metabolites
Generation of the deamidated metabolites NaMN and NaAD from the amidated precursor NR
observed by Tammell and colleagues was speculated to come from the deamidation of NMN
by the enzyme, NMN deamidase (Galeazzi et al. 2011). While NMN deamidase is a well-
known enzyme which actively participates in NAD+ metabolism in bacteria, evidence for NMN deamidase activity in mammalian cells is largely absent (Magni et al. 2008; Rongvaux et al.
2003), despite being reported in early studies (Kirchner, Watson, and Chaykin 1966; Petrack et al. 1963; Petrack et al. 1965; Tanigawa et al. 1971). It was hypothesized that the generation of the deamidated metabolites NaMN and NaAD following supplementation of NMN in the mammalian cell lines C2C12 and HEK293 is unlikely.
Consistent with this hypothesis, our study failed to detect significant production of these deamidated metabolites from exogenous NMN. Figure 3.11 A shows chromatogram peaks of
NMN/NaMN isotopologue pairs detected in C2C12 and HEK293 cells following NMN
supplementation. At time zero, cells were treated with an equivalent amount of unlabeled NMN
(due to the limited availability of labelled NMN isotope) and harvested immediately following
supplementation. The intensity of unlabeled NMN (in dark blue) detected at this time point
indicates rapid uptake of NMN into the cell. The smaller peak below this represents M+1
labelled NMN (in light blue) as well as unlabeled NaMN (M+0) which shares the same MRM
transition, that is, 336.15>124.05 for NMN (M+1) and 336.10>123.80 for NaMN (M+0). The
intensity of this smaller peak compared to the larger peak is close to the value expected from
natural abundance of M+1 labelled NMN (13.2%) (Figure 3.7 C) which suggests very little
NaMN is contributing to this peak and in turn suggests very little NaMN is generated in C2C12
and HEK293 cells following exposure to NMN. Even at the latest time point (24 hrs), this ratio
did not change significantly after cells were exposed to M+6 labelled NMN. The intensity of
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the peak representing M+6 labelled NaMN (in orange), which is also shared by M+7 labelled
NMN, is small in comparison to the intensity of the peak formed from M+6 labelled NMN (in
yellow) detected in cells. The intensities are quantified in Figure 3.11 B showing that even at
earlier time points, the level of M+6 labelled NaMN (in orange) in C2C12 (left) and HEK293
(right) cells after supplementation with M+6 labelled NMN1 (in yellow) is relatively low.
The deamidated metabolite NaAD, is synthesized from NaMN in mammals via the
enzyme NMNAT as part of the Preiss-Handler pathway (Preiss and Handler 1958b; Zhang et
al. 2003). As very little NaMN was detected in this study, the intracellular levels of NaAD in
C2C12 and HEK293 cells were also expected to be low. Supplementing C2C12 and HEK293
cells with unlabeled NMN led to an intensity of M+1 labelled NAD+ below the expected value
derived from natural abundance (26.2%) (Figure 3.8 C), suggesting very little contribution of
the unlabeled NaAD (M+0) isotopologue which shares the same MRM transition (Figure 3.12
A). The intensity of M+6 labelled NaMN following NMN1 supplementation also remained low at the late 24-hour time point in both C2C12 and HEK293 cells (in orange) compared to M+6 labelled NAD+ (in yellow), suggesting very little synthesis of NaMN in cells following
supplementation with NMN. The low levels of M+0 and M+6 labelled NaAD after
supplementation with unlabeled and M+6 labelled NMN, respectively, also suggests the high
intensity of the peak shared by M+1 labelled NAD+ and unlabeled NaAD (in light blue) most likely represents the former isotopologue, NAD+ (M+1), rather than NaAD (M+0).
Degradation of M+6 labelled NAD+ yields M+1 labelled nicotinamide, which may be salvaged to synthesize M+1 labelled NMN via NAMPT (Revollo, Grimm, and Imai 2004; Wang et al.
2006b), before final synthesis of M+1 labelled NAD+ via NMNAT (Berger et al. 2005;
Garavaglia et al. 2002; Schweiger et al. 2001; Zhang et al. 2003). The chromatogram peaks
shared by pairs of NAD+/NaAD isotopologues shown in Figure 3.12 A are quantified in Figure
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3.12 B where there is no significant detection of M+6 labelled NaAD (in orange), even at earlier
time points following supplementation with M+6 labelled NMN1.
Overall, these data show very little incorporation of the labelled moieties from M+6
labelled NMN1 into NaMN and NaAD in C2C12 and HEK293 cells. This is consistent with
the hypothesis that the appearance of deamidated metabolites observed by Trammell and
colleagues may be generated from the deamidation of NMN via NMN deamidase, an enzyme
which is present in bacteria, but not mammalian cells such as C2C12 and HEK293 used in this
study. Bacteria in the gastrointestinal tract of mammals could be contributing to the metabolism
and utilisation of orally administered NAD+ precursors such as NMN and NR, a hypothesis which is explored in the next two chapters.
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Figure 3.11 Detection of NMN/NaMN isotopologue pairs in C2C12 and HEK293 cells following supplementation with NMN. The media of C2C12 and HEK293 cells were supplemented with M+6 labelled NMN (0.1 mM) and harvested at time points 1, 2, 4, 8, and 24 hours after treatment. At time zero, cell media was supplemented with unlabelled NMN (0.1 mM) and harvested immediately. The intensity of NMN and NaMN isotopologue pairs were detected under the same peak due to shared MRM transitions. The data represents mean ± s.d. (n=2) with picomolar amounts normalised to protein concentration of each sample (pmol/mg protein). (A) Chromatogram peaks from NMN/NaMN isotopologue pairs at time zero and 24 hours after supplementation with NMN. (B) Quantification of chromatogram peaks generated from NMN/NaMN isotopologues.
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Figure 3.12 Detection of NAD+/NaAD isotopologue pairs in C2C12 and HEK293 cells following supplementation with NMN. The media of C2C12 and HEK293 cells were supplemented with M+6 labelled NMN (0.1 mM) and harvested at time points 1, 2, 4, 8, and 24 hours after treatment. At time zero, cell media was supplemented with unlabelled NMN (0.1 mM) and harvested immediately. The intensity of NAD+ and NaAD isotopologue pairs were detected under the same peak due to shared MRM transitions. The data represents mean ± s.d. (n=2) with picomolar amounts normalised to protein concentration of each sample (pmol/mg protein). (A) Chromatogram peaks generated from NAD+/NaAD isotopologues at time zero and 24 hours after supplementation with NMN. (B) Quantification of chromatogram peaks generated from NAD+/NaAD isotopologues
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3.4 Discussion
In this chapter, an LC-MS/MS method was developed to detect labelled NAD+ metabolites in
cells following supplementation with labelled isotopes of NMN. This involved optimising mass
spectrometry parameters such as the depolarisation potential (DP), collision energy (CE), cell
exit potential (CXP) and retention time (RT) for 8 different NAD+ metabolites (NMN, NR,
NAM, NAD+, NA, NaR, NaMN and NaAD). The multiple reaction monitoring (MRM) transitions for 64 different isotopologues were also pre-determined during this optimisation
process based on their respective monoisotopic masses. It was important to optimise these
parameters in order to establish the maximum signal-to-noise ratio and obtain high resolution peaks for the integration and quantification of these metabolites. These parameters will serve as an important resource for later studies in the field related to NAD+ metabolism using isotope
labelling strategies. This LC-MS/MS method was then validated using primary hepatocytes,
C2C12 and HEK293 cells as in vitro models. In doing so, the metabolic fate of NMN was
explored by tracing the incorporation of the labelled ribose and nicotinamide moieties into
NAD+ metabolites, which also provided some evidence for the mechanisms of NMN transport
into cells and its utilisation through NAD+ biosynthesis pathways.
In mammals, NMN is part of the salvage pathway where it can be converted into NAD+
in one enzymatic step via NMNAT1-3 (Berger et al. 2005; Garavaglia et al. 2002; Schweiger
et al. 2001; Zhang et al. 2003) or resynthesized from nicotinamide after NAD+ is degraded, via
NAMPT (Revollo, Grimm, and Imai 2004; Wang et al. 2006b). According to this canonical view of NAD+ metabolism in mammals, the results from this study show an increase in both
NAD+ and NAM following exogenous supplementation with NMN, as expected. Further, the
labelling of these metabolites indicated NAD+ turnover was occurring through the salvage of
nicotinamide, the by-product (along with ADP ribose) of NAD+ degradation via NAD+
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consuming enzymes (Imai et al. 2000a; Kim, Jacobson, and Jacobson 1993; Kim, Zhang, and
Kraus 2005).
One of the main interests of using labelled isotope versions of NMN was to explore the
generation of deamidated metabolites in mammalian cells following supplementation with an
amidated NAD+ precursor, as previously observed by Trammell and colleagues (Trammell et
al. 2016a). Whether this occurs through an unknown mammalian NMN deamidase or, as
previously speculated by Trammell and colleagues, through bacterial enzymes in the gut when
delivered orally, is unknown (Trammell et al. 2016a). In this study, very little labelling of
deamidated metabolites NaMN and NaAD was detected, which largely suggests that
mammalian NMN deamidase activity is unlikely, at least in primary hepatocytes and
mammalian cell lines (C2C12 and HEK293 cells) supplemented with NMN. Whether these
metabolites are generated from bacterial enzymes in the gastrointestinal tract following oral
delivery in mammals still remains a possibility and is explored in greater detail in Chapter 4
and Chapter 5.
Another highly debated topic in the field of NAD+ biology is the mechanism of NMN
transport to raise NAD+ levels inside the cell. One mechanism argues that the rapid increase of
NMN (within 1 minute) and synthesis of NAD+ (within 15 minutes in tissues) detected in cells and tissues is highly suggestive of a direct transport mechanism (Grozio et al. 2019a; Mills et
al. 2016; Yoshino et al. 2011). The elusive NMN transporter was recently suggested to come
from the family of solute carrier transporters, namely Solute Carrier Family 12 Member 8,
SLC12A8, and despite ongoing controversy in the field around the bioanalytical methods used
to determine this (Schmidt and Brenner 2019), is supposedly highly selective for NMN (Grozio
et al. 2019a). Another mechanism argues that NMN utilization by cells requires extracellular
dephosphorylation into NR before it can enter the cell (Ratajczak et al. 2016). NR then enters
through a family of equilibrative nucleoside transporters (ENTs), SLC29, comprising 4
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different types (ENT1-4) where ENT1 (SLC29A1), ENT2 (SLC29A2) and ENT3 transport
NR, while ENT4 transports adenosine (Baldwin et al. 2004). NR is then re-phosphorylated into
NMN via NR kinase enzymes, NRK1/2 (Bieganowski and Brenner 2004). Whether both
mechanisms co-exist or complement each other remains a matter of debate.
While intact labelling of NMN was observed inside the cell (Figure 3.5 A-B and
Figure 3.9 A-B), there was also an abundance of intact labelled NR detected in the in vitro models supplemented with NMN in this study (Figure 3.6 A-B, Figure 3.9 E-F). The labelled
NR pool inside the cell was determined to be higher than the labelled NMN pool (Figure 3.6
C-D) which supports the dephosphorylation of NMN to NR (Ratajczak et al. 2016). It is
important to note however, that the rapid incorporation (within first hour) of NMN detected in
C2C12 and HEK293 cells following supplementation of NMN in culture media, could provide
some evidence to support the NMN transporter mechanism in addition to the
dephosphorylation mechanism, which is a slower process reported to occur over a 24-hour time
period (Ratajczak et al. 2016; Yoshino et al. 2011). Further studies are warranted which
measure labelled NMN and NR at earlier time points (over several time points in minutes rather
than hours) to enhance our understanding of NMN transport mechanisms.
Study Limitations
One of the greatest challenges in the development of our LC-MS/MS method was the
separation of closely related pairs of NAD+ metabolites, in particular, NMN and NAD+ to their
acidic counterparts, NaMN and NaAD, respectively. Lowering the pH of the separation buffer
from pH 9 to pH 5 improved the detection of these closely related species in Chapter 4 and 5,
however this remained one of the main challenges in this study. To overcome this, the natural
abundance for the incorporation of an isotope label into NMN (13.2%) and NAD+ (26.2%) was used to assess whether unlabelled NaMN or NaAD, if present, contributed to the peak shared by M+1 labelled NMN or NAD+, respectively, following supplementation with unlabelled 117
Chapter 3 Quantifying the NAD metabolome
NMN in cells. Due to the nature of the isotope labelling strategy, where some closely related
isotopologues only differ by one mass unit, striving to achieve good chromatographic
separation is a common challenge in LC-MS based metabolomic studies (Bustamante et al.
2018). Despite this, the LC-MS/MS method developed here can detect 64 isotopologues from
8 different NAD+ metabolites following the exogenous supplementation with NMN isotopes, and was validated in three biologically relevant in vitro models which may be used to provide
future perspective on NMN transport and utilisation pathways.
Heavy labelled versions of the metabolites of interest are the ideal internal standards to
normalise sample output measures in LC-MS analysis to account for any variable loss during
sample handling and processing (Bustamante et al. 2018; Clement et al. 2019; Evans et al.
2002; Trammell and Brenner 2013). In our studies, however, the use of alternative non-heavy-
labelled internal standards, methionine sulfonate (MES), camphorsulfonic acid (CSA) and
thymine-d4 (TD4) were chosen as they were readily available, yielding consistent output values
for normalisation over multiple mass spectrometry runs and hence deemed suitable for the
investigations outlined in this thesis. They also provided an accessible option over more
expensive labelled isotope versions of compounds to study the NAD metabolome.
A sensitive component in the design of LC-MS based metabolomic studies is the time of sampling. In order to achieve the closest representation of the biological pathway being targeted, it is ideal to harvest samples rapidly to stop metabolic activity while also maintaining the stability of metabolites of interest. Recently, it was shown that metabolic activity continues to occur rapidly during and after cell harvest, and is difficult to prevent even when the extraction process is fast and performed under cold temperatures (Trefely et al. 2019). The concern is that these enzymatic activities may contribute to the artifactual detection of metabolites in whole cell extracts potentially leading to an unwanted misinterpretation of detected metabolites. One strategy to address post-harvest metabolism involves the
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introduction of a labelled substrate at the point of harvest which may be compared to samples
that were not introduced to the labelled substrate. The difference in incorporation of labelled
isotopes between these samples will be to assess the amount of post-harvest metabolism occurring in cells. This may be an important consideration for future studies as part of the data correction process in order to detect the presence and/or absence of certain metabolites with better accuracy and reliability.
Recently, it was shown that HEK293 cells grown in media supplemented with 10% fetal bovine serum (FBS) led to the degradation of nucleotides such as NMN to their corresponding nucleosides, namely NR, and in turn NR was cleaved to the corresponding bases, namely NAM (Kulikova et al. 2019). Further, this was also shown to occur even in the absence of cells, albeit to a lesser extent, indicating that FBS alone has the capacity to degrade
NAD+ intermediates. The cell culture conditions implemented in this study include
supplementation of cell culture media with 10% FBS which is the standard method for the
maintenance of growth in C2C12 and HEK293 cell lines, however this may also pose a
potential limitation in light of these FBS-degrading activities. As such, in future in vitro studies
involving supplementation with NAD+ precursors, it may be ideal to design experiments both
with and without FBS as well as with and without cells. Future studies should take into careful
consideration the influence of these culture conditions when investigating changes to NAD+
metabolites following supplementation with NAD+ precursors.
The use of inhibitors targeting enzymes involved in the synthesis or transport of NAD+
intermediates is another effective strategy which could be used in conjunction with labelled
NMN isotopes to investigate mechanisms of transport and utilisation pathways. Many of the
findings from this study were inferred purely from the detection of metabolites in samples,
however, the use of enzyme inhibitors may enhance our understanding of the pathways
involved in the utilisation and transport mechanisms associated with NMN metabolism. The
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direct utilisation of NMN for NAD+ biosynthesis in the salvage pathway is catalysed by the
enzyme, NMNAT, which can be inhibited by gallotannin (Berger et al. 2005). Mammalian
NMNATs are also responsible for the conversion of NaMN to NaAD, hence, can influence
NAD+ biosynthesis through the Preiss-Handler pathway (Berger et al. 2005; Preiss and Handler
1958b; Zhang et al. 2003). Salvage of nicotinamide, the by-product of NAD+-degradation, can
be prevented by the FK866 which inhibits NAMPT, the rate-limiting enzyme of the salvage
pathway (Revollo, Grimm, and Imai 2004; Wang et al. 2006b). To investigate the mechanism
for NMN transport through dephosphorylation into NR via the 5’-nucleotidase, CD73 (Grozio
et al. 2013), the CD73 inhibitor, adenosine-5′-(α,β-methylene) diphosphate (AOPCP), may be used (Grozio et al. 2019a). The NR transporter family of ENTs, SLC29 (ENT1-3) (Baldwin et al. 2004), which allows entry of NR into the cell can be blocked by the ENT inhibitors, S-(4-
nitrobenzyl)-6-thioinosine (NBTI) and dipyridamole (Dip), to investigate whether the
dephosphorylation of NMN to NR is required prior to its uptake and utilisation (Kulikova et
al. 2019; Ratajczak et al. 2016). The use of these inhibitors in conjunction with NMN isotopes
and LC-MS can further elucidate on the mechanisms involved in the utilisation and transport
of NMN inside the cell.
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Investigating NAD+ metabolism following supplementation with NMN in bacteria
4.1 Introduction
The microbiome plays an important role in health, encompassing all organisms that live on the
external and internal surfaces of the gastrointestinal tract (GIT), skin, oral cavity and
conjunctiva. The GIT has the highest microbial load, with estimates of up to 1014 microbial cells consisting mainly of bacteria (~98%) and a small percentage (2%) of archaea, viruses and other eukaryotes (Berg 1996; Luckey 1972; Savage 1977; Zhernakova et al. 2016). As a collective whole, the microbiome (genomes of the microbiota) contain approximately 150 times more genes than the human genome (Qin et al. 2010), encoding proteins and enzymes that play important roles in metabolic processes such as digestion and metabolism of host- indigestible foods, the immune response and breakdown of xenobiotics (Carmody and
Turnbaugh 2014; Cha et al. 2010; Fukuda et al. 2011; Hooper, Midtvedt, and Gordon 2002;
Koppel, Rekdal, and Balskus 2017; Maslowski et al. 2009; Savage 1986; Sousa et al. 2008;
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Spanogiannopoulos et al. 2016). These interactions represent a symbiotic relationship between
the host and the microbiome, which impacts metabolic processes that are important to health.
The importance of the microbiome in governing the metabolism and bioavailability of pharmaceuticals is emerging as an under-appreciated area, with drug-microbiome interactions that could impact clinical efficacy (ElRakaiby et al. 2014; Guthrie, Wolfson, and Kelly 2019;
Khalsa et al. 2017; Li and Jia 2013). While the liver has been traditionally studied as the main organ responsible for metabolising xenobiotics, recent studies emphasise the role of microbial drug metabolism in the GIT prior to interaction with the liver (Carmody and Turnbaugh 2014;
Guthrie, Wolfson, and Kelly 2019; Kang et al. 2013b; Koppel, Rekdal, and Balskus 2017;
Spanogiannopoulos et al. 2016; Sun, Chen, and Shen 2019). This is especially the case for orally delivered medicines which are not completely absorbed by the small intestines, as these compounds then encounter a far greater microbial density in the distal gut, or those which undergo metabolism through the enterohepatic circulation, where the liver and the intestines undergo the exchange of metabolites (Macfarlane and Macfarlane 2004).
Applying this phenomenon to the administration of NAD+ precursors, this study sought
to investigate how the gut microbiome alters the metabolic fate and bioavailability of orally
delivered NMN. Specifically, pathways leading to NMN incorporation by the host are poorly
understood because NMN is an atypical nutrient in our diet; nicotinic acid (NA), nicotinamide
(NAM) and nicotinamide riboside (NR) are the main constituents of vitamin B3, which the gut
microbiome and our body are accustomed to. For example, while Trammell et al. (2016)
reported a sharp spike in the appearance of the deamidated precursors nicotinic acid
mononucleotide (NaMN) and dinucleotide (NaAD) in mouse tissues following oral
administration of NR (Trammell et al. 2016a), it remains unclear how both deamidated
metabolites were raised, as mammalian deamidase enzymes are yet to be identified (Carson,
Seto, and Wasson 1987; Gazzaniga et al. 2009; Lin et al. 1972; Rongvaux et al. 2003; Shibata,
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Hayakawa, and Iwai 1986). Nonetheless, deamidation of NAD metabolites in mammalian
systems have been reported elsewhere (Kirchner, Watson, and Chaykin 1966; Petrack et al.
1963; Petrack et al. 1965; Sarma, Rajalakshmi, and Sarma 1961). These earlier findings suggest that a clear description of the pathways that actively assimilate orally delivered NMN are still missing.
One theory is that upon exposure to a bolus of amidated metabolites such as NR or
NMN, bacterial enzymes in the GIT may contribute to the generation of NaMN and NaAD via bacterial deamidation enzymes, prior to the uptake of these metabolites by the host. This is because deamidation of NAM and NMN by the deamidase enzymes PncA and PncC is regarded as a necessary enzymatic route for the utilisation of these precursors for NAD+
biosynthesis in bacteria (Galeazzi et al. 2011). If true, this could suggest a role of the
microbiome in altering the metabolism and bioavailability of exogenously administered NAD+
precursors when delivered via the oral route. These potential host-microbiome interactions in
the metabolism of NAD+ precursors could impact therapeutic strategies that involve NMN
supplementation.
4.1.1 Bacterial de novo NAD biosynthesis pathway
Bacteria such as E. coli possess enzymes capable of NAD+ biosynthesis through de novo and
salvage pathways, which include enzymatic steps that are absent in eukaryotes (Figure 4.1,
below). The NAD+ biosynthesis pathway in E. coli is well characterised. With the exception
of bacterial NAMPT (NadV), E. coli possess all the genes necessary for the de novo (nadA,
nadB, nadC, nadD, nadE, pncB) and salvage (nadR) pathways (Galeazzi et al. 2011; Kurnasov et al. 2002), including those encoding NAM and NMN deamidase enzymes pncA (Frothingham et al. 1996) and ygdA (Galeazzi et al. 2011). The deamidation of NAM and NMN by the
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deamidase enzymes PncA and PncC contribute to the utilisation of these precursors for NAD+
biosynthesis in bacteria (Galeazzi et al. 2011).
The universal precursor in the de novo NAD+ biosynthesis pathway is quinolinic acid
(QA), or quinolinate, which is essential in providing the pyridine ring for synthesis of NAD+.
In most bacteria, QA is generated from ʟ-aspartate, transported by a proton/glutamate-aspartate symporter encoded by gltP gene (Tolner et al. 1992). First, the enzyme NadB (ʟ-aspartate oxidase, EC 1.4.3.16, encoded by nadB) oxidises ʟ-aspartate forming the intermediate α- iminoaspartate in the cytoplasm (Flachmann et al. 1988; Nasu, Wicks, and Gholson 1982;
Seifert et al. 1990). Second, the enzyme NadA (quinolinate synthase, EC 2.5.1.72, encoded by nadA) catalyses the formation of QA from α-iminoaspartate in the presence of dihydroxyacetone phosphate (DHAP) in the cytoplasm (Ceciliani et al. 2000; Flachmann et al.
1988), thus completing the aspartate-to-quinolinate pathway.
NAD+ is then synthesised from QA via the Preiss-Handler pathway (Preiss and Handler
1958a, 1958b). First, QA undergoes phosphoribosylation and decarboxylation by NadC
(quinolinic acid phosphoribosyl transferase, EC 2.4.2.19, encoded by nadC) to form NaMN
(Bhatia and Calvo 1996; Gholson et al. 1964). NaMN can also be regenerated from another precursor, nicotinic acid (NA) by the enzyme PncB (nicotinic acid phosphoribosyltransferase,
EC 6.3.4.21, encoded by pncB) in the presence of phosphoribosyl pyrophosphate (PRPP) and
ATP (Imsande 1961). NA may be taken up by niacin transporters NiaX (Johnson et al. 2015;
Rodionov et al. 2008). NaMN is then adenylated into NaAD by the nicotinic acid mononucleotide adenylyltransferase enzyme NadD (EC 2.7.7.18, encoded by nadD) (Imsande
1961; Mehl, Kinsland, and Begley 2000; Zhang et al. 2002).
In the last step of bacterial NAD+ biosynthesis, NaAD undergoes amidation by the NAD synthetase NadE (EC 6.3.1.5, encoded by nadE) into NAD (Spencer and Preiss 1967). The
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amidation of NaAD is dependent on free ammonia (NH3) as a source of the amide in most
prokaryotes, including E. coli (De Ingeniis et al. 2012; Spencer and Preiss 1967).
4.1.2 NAD+ is salvaged by deamidation in bacteria
It is important to note that NMN is not an intermediate of the de novo pathway. It is instead
part of the salvage pathway, used to regenerate NAD+ following its hydrolysis by NAD+
consuming enzymes, releasing free NAM, which can be recycled into NAD+ via NMN
(Andreoli et al. 1972; Olivera and Lehman 1967). NAM may enter through niacin transporters
NiaX (Johnson et al. 2015). Interestingly, in bacteria, NAM or NMN are utilised by first
converting them into their acid forms, NA and NaMN respectively by the nicotinamide deamidase PncA (EC 3.5.1.19, encoded by pncA) (Frothingham et al. 1996) and the NMN
deamidase PncC (EC 3.5.1.42, encoded by ygaD) (Foster and Baskowsky-Foster 1980;
Galeazzi et al. 2011; Imai 1973; Manlapaz-Fernandez and Olivera 1973).
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Figure 4.1 NAD+ biosynthesis pathways in bacteria. The synthesis of NAD+ in bacteria can occur via three pathways. The de novo synthesis pathway initiates with the metabolism of L-aspartate in most bacteria, but L-tryptophan in some groups such as those in the taxonomic order Xanthomonodales, Flavobacteriales and Burkholderiales. Both L-tryptophan and L-aspartate ultimately form quinolinic acid (QA) which is converted into nicotinic acid mononucleotide (NaMN) via NadC in the Preiss- Handler pathway. NaMN may also be synthesised from the precursor nicotinic acid (NA) via PncB. NaMN is then adenylated into nicotinic acid adenine dinucleotide (NaAD) via NadD followed by amidation into NAD+ via NadE. In the salvage pathway, nicotinamide mononucleotide (NMN) may enter the periplasmic space via the porin OmpP2, where it may be metabolised by two separate routes to form NAD+. It may either be hydrolysed to nicotinamide (NAM) and a ribose moiety via NMN glycohydrolase or undergo dephosphorylation into nicotinamide riboside (NR) via periplasmic 5’-nucleotidase UshA. NAM may enter via the niacin transporters NiaX while NR may enter through PnuC where the latter is re-phosphorylated into NMN via the NR kinase activity of NadR. NAM and NMN may both undergo deamidation via deamidases PncA and PncC to form NA and NaMN respectively and synthesise NAD+ via the Preiss-Handler pathway. The nicotinamide mononucleotide adenylyltransferase (NMNAT) activity of NadR is weak in most bacteria, hence relying mostly on NMN deamidation for NAD+ biosynthesis. In some species such as Haemophilus influenzae however, NAD+ biosynthesis from NR via the activity of trifunctional NadR is essential as they lack genes that encode for de novo pathway and Preiss- Handler pathway enzymes.
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There are two reasons for the deamidation of NAM and NMN. First, NMN can inhibit
NAD+-dependent DNA ligation in bacteria (Chen et al. 2002; Geider 1972; Zimmerman and
Oshinsky 1969). Intracellular NMN builds up quickly during the exponential growth of bacteria
due increased rate of DNA repair imposing a greater conversion of NAD+ into NMN (Lehnman
1974; Olivera and Lehman 1967). Deamidation therefore acts to promote growth and repair by
reducing the availability of NMN, which inhibits DNA ligase. NMN is salvaged through the
NMN deamidase PncC, yielding NaMN as a substrate for NAD+ synthesis by the Preiss-
Handler pathway (Galeazzi et al. 2011).
The second reason for NMN deamidation in bacteria is that NaMN is the preferred phosphoribosyl donor for the enzyme cobaltochelatase (CobT) (Agerholm et al. 2017; Warren et al. 2002). This enzyme catalyses the last step of adenosylcobalamin (vitamin B12) biosynthesis, an important cofactor involved in DNA synthesis, red blood cell production and various functions of the nervous system (Fenech 2001; Reynolds 2006). Even in bacterial species that are capable of NAM and NMN salvage via the non-deamidated route, such as in
E. coli, the deamidation route is preferred. These processes emphasise why deamidation to
NAD+ synthesis are preferred in bacteria, as the amidated intermediate NMN can impair DNA
ligase, and the deamidated intermediate NaMN is an important precursor for cofactor synthesis
(Maggio-Hall and Escalante-Semerena 2003).
4.1.3 Potential NMN uptake routes in bacteria
Gram-negative bacteria such as E. coli have a bi-dermal membrane consisting of an inner and outer membrane separated by an area known as the periplasmic space. NMN enters the periplasmic space through a porin (channel-forming protein) known as OmpP2 (Andersen et
al. 2003; Pullen et al. 1995). Here, the assimilation of NMN into the cytoplasm can occur via
two different routes.
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The first route involves the dephosphorylation of NMN to nicotinamide riboside (NR)
by the 5’nucleotidase UshA (EC 3.1.3.5, encoded by ushA) (Burns and Beacham 1986).
Transport of NR across the inner membrane involves the transporter PnuC (Jaehme, Guskov,
and Slotboom 2014) and once in the cytoplasm, NR is then re-phosphorylated into NMN by
NadR (NR kinase, EC 2.7.7.1, encoded by nadR).
The second route involves degradation of NMN by the enzyme, NMN glycohydrolase
(EC 3.2.2.14), which is embedded in the inner membrane, forming NAM and ribose 5- phosphate (Andreoli et al. 1972). As NAM is a relatively small molecule, it can permeate the inner and outer membranes, entering the cytoplasm where it may undergo deamidation through
PncA, forming NA. As in de novo biosynthesis from aspartate, NA is used as a substrate for
NAD+ synthesis by the Preiss-Handler pathway enzymes, PncB (NA to NaMN), NadD (NaMN
to NaAD) and NadE (NaAD to NAD+).
Interestingly, NadR is a uniquely tri-functional enzyme which can carry out the following
roles:
1. Transcriptional downregulation of the genes pncB (salvage pathway), nadA and nadB
(de novo pathway) when NAD+ levels in the cell are replete,
2. Interaction with the NR transporter PnuC to promote the uptake of NR into the cell
when NAD+ levels are scarce,
3. Dual NR kinase (NRK) and NMNAT enzymatic activity for the substrates NR and
NMN, respectively (Foster and Baskowsky-Foster 1980; Grose, Bergthorsson, and
Roth 2005; Penfound and Foster 1999; Raffaelli et al. 1999).
NadR is therefore an important regulatory enzyme for coordinating the utilisation and
transport of NMN and NR through both transcriptional and direct enzymatic control. NadR
activity is essential for some species of bacteria, such as Haemophilus influenzae, that lack
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genes that encode for the enzymes involved in de novo NAD+ biosynthesis (Singh et al. 2002).
In most other bacteria however, NadR has weak NMNAT activity and its activity is insufficient
to prevent the accumulation of NMN, and cells instead rely on PncC to for this activity (Grose,
Bergthorsson, and Roth 2005).
4.1.4 Key differences compared to eukaryotic pathway
There are three main features which bacterial and eukaryotic pathways differ. First, de novo synthesis in higher eukaryotes starts with ʟ-tryptophan to derive the pyridine ring of QA (refer to Chapter 1, Section 2.6.1), instead of condensing ʟ-aspartate. Certain taxonomic groups, such
as Xanthomonadales (Order), Flavobacteriales (Order) and Burkholeriales (Order) do not
express genes encoding nadA and nadB, and therefore rely on QA synthesis from ʟ-tryptophan
(Kurnasov et al. 2003; Lima, Varani, and Menck 2009) which can be transported into the cell
via three transporters, Mtr, TnaB and AroP (Yanofsky, Horn, and Gollnick 1991), however,
most bacterial species rely on QA synthesis from ʟ-aspartate.
Importantly, the deamidation of NAM and NMN which occurs in bacteria does not
occur in mammals. In eukaryotes, NAM is converted to NMN by the nicotinamide
phosphoribosyltransferase NAMPT (EC 2.4.2.12, encoded by NAMPT) (Preiss and Handler
1957). NMN is then used as a substrate by nicotinamide mononucleotide adenylyltransferase
enzymes (NMNAT, EC 2.7.7.1 encoded by Nmnat) to generate the final step of NAD+
(Emanuelli et al. 2001; Raffaelli et al. 2002; Zhang et al. 2003). Phylogenetic analysis of
bacterial genomes reveals an absence of NAMPT (NadV) and NMNAT (NadM and NadR)
(Galeazzi et al. 2011), and bacteria must instead rely on PncA and PncC to convert the amidated
metabolites nicotinamide and NMN to their acid forms of nicotinic acid and NaMN before they
become substrates for NAD+ biosynthesis. In eukaryotes the canonical pathway for NAD+
biosynthesis from NR involves its phosphorylation into NMN via NR kinase (NRK1/2)
(Bieganowski and Brenner 2004), prior to direct conversion of NMN into NAD+ via NMNAT
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1-3 (Raffaelli et al. 2002; Schweiger et al. 2001; Zhang et al. 2003), a pathway which clearly
does not involve deamidated intermediates.
Another difference in NAD metabolism between bacteria and eukaryotes is that
amidation in eukaryotes is fuelled by glutamine (C5H10N2O3) via the enzyme NAD synthetase
(NADSYN1) (Hara et al. 2003; Preiss and Handler 1958a, 1958b), whereas the bacteria use
free ammonia as a course of nitrogen for amidation of NaAD into NAD+. This likely reflects
differences in the availability of these nitrogen sources.
The aim of this work was to examine how bacteria metabolise NMN, in particular the
potential conversion of NMN into deamidated products. First, we reviewed the literature to
build the bacterial NAD pathways to identify functional contributions that are distinguished
from mammalian counterparts. We then generated metabolomic data from an in vitro model,
in which NMN was supplemented into the broth of cultured E. coli bacteria. The reconstructed
pathways and the metabolite data were then integrated to resolve NAD pathways that are active
in E. coli. Most importantly, the detection of labelled intermediates in the culture supernatant
upon supplementing labelled NMN indicated a putative role of the commensal gut microbiome in host NAD+ homeostasis. Following the work described in this chapter, an in vivo mouse
model is described in the next chapter to identify a host-microbiome integrated model of NAD+
homeostasis.
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4.2 Methods
4.2.1 Bacterial culture and NMN treatment
A stab culture of the E. coli strain OP50 was inoculated into sterile Luria-Bertani (LB) broth
(consisting of 10 g/L tryptone, 5 g/L yeast extract and 10 g/L sodium chloride in deionized water) under aseptic conditions and incubated overnight at 37℃ on a shaking platform set at
200 rpm. This strain of E. coli (OP50) is a standard food source for growing the model nematode organism Caenorhabditis elegans (C. elegans), commonly used as a robust genetic
model for lifespan studies.
To measure the growth rate of E. coli, the overnight culture was sub-cultured (1:200)
into sterile LB broth in a new flask and the optical density was measured at 600 nm (OD600)
every 20 minutes (approximate doubling time) (Figure 4.1 A). This growth curve was used to
ensure samples were collected during the early-mid exponential growth phase (OD600 < 0.70),
as bacterial enzymes are more active during exponential growth phase than stationary phase
(Rahman et al. 2006). For samples, the overnight culture was sub-cultured (1:200) and
aliquoted into smaller volumes. The cultures were then supplemented with either vehicle
(water) or unlabelled NMN (0.5 mM) and collected at the following time points; time zero
(before NMN), time zero (immediately after NMN), 120, 160 and 180 minutes. The supernatant
of cells was separated from the cells via centrifugation (5000g for 10 minutes at 4℃) and stored immediately at -30℃. Meanwhile, the cell pellet was resuspended in cold (4°C) saline solution
(0.9% NaCl) and centrifuged as above, to rinse away residual media before storage at -30℃.
The OD600 (Figure 4.1 B) was measured for each sample and used to normalise metabolite
levels after LC-MS/MS analysis. In a separate experiment, the growth rate of E. coli (Figure
4.2 A) and OD600 from samples supplemented with M+6 labelled NMN1 (Figure 4.2 B) was
also measured at time zero (before NMN), time zero (after NMN), and 140, 160 and 180
minutes after supplementation with NMN1 at a final concentration of 0.1 mM. The reason for
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Chapter 4 Investigating microbial NAD+ metabolism using isotope labelled NMN at a concentration of 0.1 mM, compared to initial experiments with unlabelled NMN at 0.5 mM, was the limited availability of isotope labelled material.
Figure 4.2 E. coli growth curve and sample OD600 values. (A) E. coli (OP50) bacteria incubated overnight was sub-cultured (1:200) in sterile LB broth. The OD600 was measured every 20 minutes to generate a growth curve. (B) E. coli culture broth was supplemented with 0.5 mM NMN and collected at time points 0 (before NMN), 0 (after NMN), 120, 160 and 180 minutes. OD600 was measured for samples at each collection time point. Data represents mean ± s.d. (n=3 technical replicates for control, 5 technical replicates for NMN, each sample OD600 read in triplicate).
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Figure 4.3 E. coli growth curve and sample OD600 values. (A) E. coli (OP50) bacteria incubated overnight was diluted 1:200 with fresh LB broth. The OD600 was measured every 20 minutes to generate a growth curve. (B) E. coli culture broth was supplemented with 0.1 mM NMN1 and collected at time points 0 (before NMN1), 0 (after NMN1), 140, 160 and 180 minutes. OD600 was measured for samples at each collection time point. Data represents mean ± s.d. (n=1 technical replicates for control, 3 technical replicates for NMN1, each sample OD600 read in triplicate).
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4.2.2 Extracting NAD+ metabolites from E. coli cell lysates and culture supernatants.
To separate the supernatant from cells, samples were immediately centrifuged at 5000 g for 10 minutes at 4℃. Samples were collected at 0, 120, 160 and 180 minutes following supplementation with unlabelled NMN. These time points represent the early-mid exponential phase of growth, determined by the first, second and third doubling time points measured by
OD600. When treating with labelled NMN1, samples were collected at time points 0 (before
and after the addition of NMN1), 140, 160 and 180 minutes. Following centrifugation, the
supernatant was collected and stored at -30℃ until ready for further processing. The remaining
cell pellet (cell lysate) was rinsed once with a cold saline (0.9% NaCl) solution to remove
residual media by centrifugation at 5000 g for 10 minutes at 4℃. The supernatant was
discarded, and the final pellet was snap frozen in liquid nitrogen and stored at -30℃ until ready
for further processing.
To extract NAD+ metabolites from cell lysates, pellets were resuspended in an extraction buffer (500 µL of a 2:2:1 mixture of acetonitrile, methanol and water at -30℃) by
repeated pipetting. These resuspended cells were centrifuged at 16,000 g for 10 minutes at 4℃
and supernatant, containing the extracted NAD+ metabolites, was transferred into a new tube.
Using a centrifugal vacuum system (Savant SpeedVac SPD140DDA Vacuum Concentrator,
Thermo Fisher Scientific), in the solution was evaporated to yield a dried pellet, which was
resuspended in LC-MS-grade water (50 µL, Fisher Healthcare, cat.no. PI51140) before
centrifugation at 16,000g for 10 minutes at 4℃. The supernatant was transferred into HPLC
vials and promptly analysed by LC-MS/MS (see Chapter 2 Section 2.3 LC-MS/MS). To extract
NAD+ metabolites from the supernatant, 30 µL of media was combined with 120 µL of
extraction buffer, consisting of a 1:1 ratio of acetonitrile and ethanol (HPLC grade) before
centrifugation at 16,000g for 10 minutes at 4℃. The supernatant was transferred to a new tube,
snap frozen in liquid nitrogen and stored at -30℃ before further processing. The supernatant
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was dried down as above using the speed vac system and the dried pellet was resuspended in
30 µL of LC-MS-grade water, centrifuged as above and transferred into HPLC vials for LC-
MS/MS analysis. The internal standards, CSA, MES and thymine-d4 (see Chapter 2, Section
2.2.1 Preparation of extraction buffer) were added to each respective extraction buffer prior to extraction. The internal standard thymine-d4 was used for normalisation of peak intensities before normalising samples to the OD600 value.
Table 4.1 (below) details mass spectrometry (MS) parameters including DP, CE, CXP
and RT, and MRM transitions used to select and identify NAD+ metabolites. As this experiment
was repeated and analysed as two separate batch experiments (Run 1 and Run 2), the MS
parameters are shown for each LC-MS analysis. The first batch (Run 1) included one control
(water) and three NMN (0.5 mM) treated samples whereas the second batch (Run 2) included
two control and two NMN treated samples. Overall, the data represent three control and five
NMN treated samples. Similarly, Table 4.2 and Table 4.3 (below) shows the MS parameters
and MRM transitions used to select and identify NAD+ isotopologues from amidated (Table
4.2) and deamidated (Table 4.3) metabolites which have incorporated isotopes from labelled
ribose and nicotinamide moieties following supplementation with NMN1 (0.1 mM) and
includes one control and three NMN1 treated samples. The MS parameters, DP, CE and CXP
remained the same throughout all experiments and the slight changes in retention times are
likely due to differences in laboratory conditions and LC-column status.
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Table 4.1 MRM transitions and mass spectrometry (MS) parameters to detect NAD+ metabolites in E. coli (supernatant and lysates) following supplementation with unlabelled NMN using LC-MS/MS. Run 1 included one control and three NMN treated samples whereas Run 2 included two control and two NMN treated samples.
Run 1 MRM MS parameters Supernatant Lysate Metabolite Q1 Q3 DP CE CXP RT RT Isotopologue Q1_Q3 label (m/z) (m/z) (V) (V) (V) (mins) (mins)
NAM_0_0 NAM (M+0) 122.8 79.8 80.0 30.0 25.0 2.0 2.0 NMN_0_0 NMN (M+0) 335.2 123.1 47.7 24.0 11.0 18.1 18.1 NR_0_0 NR (M+0) 255.0 123.0 64.3 29.6 12.6 11.2 11.2 NAD+_0_0 NAD+ (M+0) 664.0 428.1 33.0 36.0 31.0 16.9 16.9 NA_0_0 NA (M+0) 123.8 77.8 70.0 25.0 10.0 4.2 4.4 NaR_0_0 NaR (M+0) 256.0 124.0 46.0 18.3 13.3 10.1 10.1 NaMN_0_0 NaMN (M+0) 336.1 123.8 66.0 27.7 10.7 17.8 17.8 NaAD_0_0 NaAD (M+0) 665.0 428.1 139.0 35.0 38.0 17.2 17.2 Run 2 MRM MS parameters Supernatant Lysate Metabolite Q1 Q3 DP CE CXP RT RT Isotopologue Q1_Q3 label (m/z) (m/z) (V) (V) (V) (mins) (mins) NAM_0_0 NAM (M+0) 122.8 79.8 80.0 30.0 25.0 2.0 2.0 NMN_0_0 NMN (M+0) 335.2 123.1 47.7 24.0 11.0 18.1 18.4 NR_0_0 NR (M+0) 255.0 123.0 64.3 29.6 12.6 11.2 11.2 NAD+_0_0 NAD+ (M+0) 664.0 428.1 33.0 36.0 31.0 16.9 16.9 NA_0_0 NA (M+0) 123.8 77.8 70.0 25.0 10.0 4.6 4.4 NaR_0_0 NaR (M+0) 256.0 124.0 46.0 18.3 13.3 10.1 10.1 NaMN_0_0 NaMN (M+0) 336.1 123.8 66.0 27.7 10.7 17.8 17.6 NaAD_0_0 NaAD (M+0) 665.0 428.1 139.0 35.0 38.0 17.2 17.2
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Table 4.2 MRM transitions and MS parameters to detect NAD+ isotopologues of amidated metabolites in E. coli (supernatant and lysates) following supplementation with NMN1 using LC-MS/MS.
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Table 4.3 MRM transitions and MS parameters to detect NAD+ isotopologues of deamidated metabolites in E. coli (supernatant and lysates) following supplementation with NMN1 using LC-MS/MS.
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4.3 Results
4.3.1 Supplementation of NMN in E. coli – changes to NAD+ metabolites.
To investigate the incorporation of NMN into the NAD+ metabolome in bacteria, E. coli were cultured in the presence of labelled and unlabelled NMN. Exogenous supplementation of NMN in E. coli led to changes in both amidated (NMN, NR, NAM and NAD+) and deamidated (NA,
NaR, NaMN and NaAD) metabolites, as expected. This was further supported by detecting the incorporation of labelled ribose (13C) and nicotinamide (15N) moieties into both the cell lysate and supernatant following supplementation with M+6 labelled NMN1 (Figure 4.5 and Figure
4.7). A breakdown of these findings will be discussed in detail in the following paragraphs.
NMN in the supernatant was detected in abundance immediately following its addition
(Figure 4.4 A), but rapidly diminished at subsequent time points indicating almost instant assimilation (Figure 4.4 B) and subsequent conversion into downstream metabolites during the exponential growth phase. This was confirmed by detection of M+6 labelled NMN in both the supernatant (Figure 4.5 A) and cell lysate (Figure 4.5 B), with a similar pattern of detection across the time points following supplementation with NMN1.
NMN can also enter the periplasmic space through the OmpP2 porin (channel-forming protein) (Andersen et al. 2003), where it undergoes dephosphorylation via the periplasmic
5’nucleotidase UshA (Burns and Beacham 1986; Wang et al. 2014b) to form NR. Consistent with this, NR was detected in the supernatant at all time points (Figure 4.4 C) which was confirmed through labelling at both the ribose and nicotinamide moieties (Figure 4.5 C). As
UshA is a periplasmic enzyme, this may be the result of contamination of periplasmic metabolites such as NR during the separation of the supernatant from cell lysate via centrifugation. NR may then betransported across the inner cytoplasmic membrane via PnuC
(Jaehme, Guskov, and Slotboom 2014), where it is re-phosphorylated by NadR in the cytoplasm to generate NMN. Interestingly, very little NR could be detected in the cell lysate
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(Figure 4.4 D), including both labelled and un-labelled versions (Figure 4.5 D). This could be attributed to the high catalytic efficiency of NadR reported in previous studies (Kurnasov et al.
2002), preventing NR efflux back through PnuC and trapping NMN inside the cell to encourage
NAD+ biosynthesis (Grose, Bergthorsson, and Roth 2005).
In addition to detecting NR in the supernatant, an abundance of NAM was also observed
in the supernatant (Figure 4.4 E), but very little was detected in the cell lysate (Figure 4.4 F).
This was confirmed again through M+1 label at the pyridine ring base of NAM in Figure 4.5
E (supernatant) and Figure 4.5 F (cell lysate). NAM may enter the cytoplasm through niacin
transporters NiaX (Johnson et al. 2015) and NAM can be generated from the degradation of
NMN by NMN glycohydrolase embedded in the inner membrane. Once in the cytosol, NAM
can serve as the primary precursor for NA through its deamidation via PncA.
Consistent with this, treatment with unlabelled NMN increased NA in cell lysates
(Figure 4.6 B), and this was confirmed by the incorporation of labelled NMN into M+1
labelled NA (Figure 4.7 B). Interestingly, a small amount of NA was also detected in the
supernatant (Figure 4.6 A and Figure 4.7 A for labelled NA) almost immediately after
supplementation with NMN. This could suggest that some NA in the cell lysate permeates out
of the inner and outer membrane and into the supernatant, where in the context of an integrated
host-microbiome model of NAD homeostasis, it could be available as a precursor for cellular
uptake in mammals.
NaMN in the cell lysate was detected following supplementation with unlabelled NMN
(Figure 4.6 F) and confirmed through M+6 labelled NaMN following supplementation with
labelled NMN1 (Figure 4.7 F). NaMN is a substrate for NaAD synthesis via NadD, however
no NaAD was detected in the cell lysate (Figure 4.6 H) following supplementation with NMN,
even following supplementation with labelled NMN1 (Figure 4.7 H). One possibility for this
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is that NaAD is rapidly assimilated into NAD+ during the exponential growth phase of bacteria, circumventing its detection. Nonetheless, due to the loss of 15N label by deamidation, the incorporation of NMN into NAD via NaAD can be inferred based on the detection of M+6 labelled NAD+ (Figure 4.5 H) in the cell lysate following NMN1 supplementation.
Both NAD+ (Figure 4.4 G) and NaAD (Figure 4.6 G) were largely absent from the
supernatant, which was expected due to the absence of dedicated transporters and the large- highly charged structures that are impermeable to membrane diffusion. This was mirrored in cells treated with labelled NMN1, as M+6 labelled NAD+ in the supernatant (Figure 4.5 G)
and NaAD (Figure 4.7 G) was not detected. In contrast, NaR, a molecule which was not
expected to be present in the extracellular space, was detected in the supernatant (Figure 4.6
C) and largely absent in the cell lysate (Figure 4.6 D). In human cells, while NaR may be
generated from the dephosphorylation of NaMN via cytosolic nucleotidases II (CN-II) and III
(CN-III) (Kulikova et al. 2015; Nikiforov et al. 2011), NaR was detected exclusively in the
supernatant, not in the cell lysate in this study, for reasons that are puzzling. Even if there were
bacterial homologues for the CNII and CNIII found in humans, bacterial 5’-nucleotidases
found in the cytosol seem to be conditionally lethal (Innes, Beacham, and Burns 2001). Further,
whether there is a specific mechanism for NaR to be transported from the cytosol into the
extracellular compartment remains unclear. As a nucleoside, NaR may be transported across
the plasma membrane through the equilibrative nucleoside transporters (ENTs) SLC29A1 and
SLC29A2 (Baldwin et al. 2004; Kulikova et al. 2015), however transport of NaR across the
plasma membrane through ENTs is reportedly less efficient than the transport of NR, the
amidated counterpart of NaR (Belenky et al. 2009). To test whether the presence of NaR in
the supernatant is due to increased transport of this metabolite across the plasma membrane
through ENTs, the pharmacological inhibitors of nucleoside transporters, S-(4-nitrobenzyl)-6-
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Chapter 4 Investigating microbial NAD+ metabolism thioinosine (NBTI) and dipyridamole (Dip) may be used to further elucidate on these mechanisms in future studies (Kulikova et al. 2019; Nikiforov et al. 2011).
NaMN was also detected in the supernatant of cells (Figure 4.6 E), however enzymes that generate NaMN including the NMN deamidase PncC, the NA phosphoribosyltransferase
PncB, and NaR kinase are all located in the cytosol. Further, NaMN is not among the metabolites known to freely permeate the plasma membrane, nor has a specific transporter been identified for NaMN-specific efflux from the cell in any species. Further investigation is required to determine how NMN treatment can lead to the formation of NaMN in the supernatant in bacteria. One possibility is that they are released into the supernatant following their synthesis to facilitate NAD+ biosynthesis in neighbouring cells, a mechanism which has been previously reported in HEK293 (human embryonic kidney) and HepG2 (hepatocellular carcinoma) cell lines (Kulikova et al. 2015). In the context of a putative microbiome-host interaction, this may support the growth of bacterial populations in the GIT and/or serve as alternative precursors for uptake and utilisation by host cells.
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Figure 4.4 Detection of amidated NAD+ metabolites in the supernatant and cell lysate from E. coli following supplementation with NMN. E. coli bacteria grown in suspension were supplemented with unlabelled NMN (0.5 mM). Whole cultures (supernatant + cells) were collected at time point, 0 (before NMN), 0 (after NMN), 120, 160 and 180 minutes after NMN supplementation. Following the separation of the supernatant from the cell lysate, NAD+ metabolites were extracted and detected using LC-MS/MS. Data represent mean ± s.d (n=3-5). (A) NMN, (C) NR, (E) NAM and (G) NAD+ in the supernatant. (B) NMN (D) NR, (F) NAM and (H) NAD+ in the cell lysate.
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Figure 4.5 Detection of amidated NAD+ metabolites in the supernatant and cell lysate from E. coli following supplementation with NMN1. E. coli bacteria grown in suspension were supplemented with M+6 labelled NMN1 (0.1 mM). Whole cultures (supernatant and cells) were collected at time points, 0 (before NMN1), 0 (after NMN1), 140, 160 and 180 minutes after NMN1 supplementation. Following separation of the supernatant from the cell lysate, NAD+ metabolites were extracted and detected using LC-MS/MS. Data represent mean ± s.d (n=3-5). (A) NMN (M+6), (C) NR (M+6), (E) NAM (M+1) and (G) NAD+ (M+6) in the supernatant. (B) NMN (M+6) (D) NR (M+6), (F) NAM (M+1) and (H) NAD+ (M+6) in the cell lysate.
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Figure 4.6 Detection of deamidated NAD+ metabolites in the supernatant and cell lysate from E. coli following supplementation with NMN. E. coli bacteria grown in suspension were supplemented with unlabelled NMN (0.5 mM). Whole cultures (supernatant and cells) were collected at time points, 0 (before NMN), 0 (after NMN), 120, 160 and 180 minutes after NMN. Following separation of the supernatant from the cell lysate, NAD+ metabolites were extracted and detected using LC-MS/MS. Data represent mean ± s.d (n=3-5). (A) NA, (C) NaR, (E) NaMN and (G) NaAD in the supernatant. (B) NA, (D) NaR, (F) NaMN and (H) NaAD in the cell lysate.
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Figure 4.7 Detection of deamidated NAD+ metabolites in the supernatant and cell lysate from E. coli following supplementation with NMN1. E. coli bacteria grown in suspension were supplemented with M+6 labelled NMN1 (0.1 mM). Whole cultures (supernatant and cells) were collected at time points, 0 (before NMN1), 0 (after NMN1), 140, 160 and 180 minutes after NMN1. Following separation of the supernatant from the cell lysate, NAD+ metabolites were extracted and detected using LC- MS/MS. Data represent mean ± s.d (n=3-5). (A) NA (M+1), (C) NaR (M+6), (E) NaMN (M+6) and (G) NaAD (M+6) in the supernatant. (B) NA (M+1), (D) NaR (M+6), (F) NaMN (M+6) and (H) NaAD (M+6) in the cell lysate.
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4.4 Discussion
NAD+ plays a critical role as a redox cofactor in metabolic reactions that are conserved across
bacteria and higher eukaryotes (Gazzaniga et al. 2009). As such, NAD+ precursors
administered through the oral route could benefit both the microbiome and the mammalian
host. This study therefore sought to shed light on pathways that may be active in the
microbiome, and could contribute to an integrated model of NAD+ homeostasis. Trammell et
al. (2016) reported the detection of NaMN and NaAD in mammals following the administration
of NR, despite the absence of a canonical pathway for NR deamidation in mammals. Our
results suggest that contributions from the gut microbiome in deamidating NMN could
contribute to this striking phenomenon. Based on our results, we proposed that NR is
phosphorylated into NMN and then deamidated into NaMN (Galeazzi et al. 2011; Sorci et al.
2014), which is converted into NaAD and then NAD+ (Preiss and Handler 1958a, 1958b).
We detected NaMN in the cell lysate following treatment with NMN, though could not
detect NaAD. This indicated that NMN deamidation was active, presumably by the bacterial
NMN deamidase PncC. Despite the inability to detect NaAD as an intermediate, the production
of M+6 labelled NAD+ at later timepoints confirmed that NMN did undergo incorporation via
deamidation. Our inference draws upon previous work that in bacteria, NAD+ must be synthesised from NaAD; further work would be required to validate these findings. The direct utilisation of NMN into NAD+ via NadR is unlikely, although may explain the absence of
NaAD in the cell lysate. NadR has both NR kinase as well as NMN adenylyltransferase
(NMNAT) activity, however previous studies show that the latter activity is weak and physiologically irrelevant (Grose, Bergthorsson, and Roth 2005). The fact that NMN utilisation
in bacteria has been reported to occur mostly through its deamidation into NaMN, rather than
through the direct synthesis of NAD+, supports this (Kurnasov et al. 2002). Another possible
explanation for the absence of NaAD in the cell lysate could be of its rapid assimilation into
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NAD+ by the enzyme NadE, given abundant availability of ammonia as a nitrogen donor for
NaAD amidation into NAD (Willison and Tissot 1994).
The labelling of the amidated metabolites NMN, NR, and NAM as well as the deamidated metabolites, NA, NaR and NaMN contributed to resolving how NMN is taken up and utilised in E. coli. (Figure 4.1). The detection of a substantial pool of NR in the supernatant strongly suggests NMN was rapidly dephosphorylated to NR by the 5’-nucelotidase UshA
(Burns and Beacham 1986) in the periplasmic space, following which it can then enter the cytoplasm through the nucleoside transporter PnuC (Jaehme, Guskov, and Slotboom 2014).
Labelled NMN and NR in the cell lysate were detected immediately after supplementation with
NMN1, and both rapidly diminished at later time points (Figure 4.4 A and C, Figure 4.5 A and C). This suggests they were rapidly assimilated into the NAD metabolome.
Our results then suggest a potential bifurcation at NMN. The first is a direct intracellular route involving rapid conversion of NR-derived NMN to NaMN; this is consistent with the high catalytic efficiency (Kcat/Km) of NadR and PncC (Galeazzi et al. 2011; Kurnasov et al.
2002). The second extracellular route involves NMN hydrolysed into NAM by NMN glycohydrolase, an enzyme embedded in the inner membrane facing the periplasmic side
(Andreoli et al. 1972), followed by deamidation of NAM into NA by PncA (Frothingham et al.
1996). This was inferred based on the detection of labelled NAM and NA in the supernatant.
NAM diffuses across the inner membrane, and NA is actively transported via NiaP (Jaehme and Slotboom 2015; Jeanguenin et al. 2012), before both finally converging at NaMN via PncB
(Wubbolts et al. 1990).
The detection of NaR and NaMN in the supernatant was unexpected. It will be important to follow these results up to determine whether this is an experimental artefact from necrotic and lysed cells or if it represents the true transport of these metabolites from bacteria.
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The NMN deamidase PncC is located in the cytoplasm, and there is no identified transporter
for NaMN. NaR in the supernatant could be generated from the dephosphorylation of NaMN
by 5’-nucleotidases in the cytosol, which have been identified in yeast as Isn1 and Std1 (Bogan et al. 2009) and humans cells as CNII and CNIII (Kulikova et al. 2015), though homologues
have not been identified in bacteria. It is not known whether the 5’-nucleotidase UshA,
responsible for the dephosphorylation of NMN to NR, can also dephosphorylate NaMN to
NaR, or whether any other 5’-nucleotidase is responsible for dephosphorylating NaMN in
bacteria, which may be located in different cell compartments (Bogan and Brenner 2010).
Importantly, the absence of NaR in cell lysates is consistent with the fact that 5’-nucleotidase activity in the cytoplasm is lethal (Innes, Beacham, and Burns 2001) and that NaR is also poorly imported (Belenky et al. 2009).
In summary, data from this study largely support canonical routes of NMN uptake and utilisation in bacteria through both dephosphorylation into NR and hydrolysis to release NAM.
The generation of the deamidated metabolite NaMN was also observed, supporting the hypothesis that bacterial enzymes in the GIT may be contributing to the metabolism of orally administered NAD+ precursors such as NMN. Although the deamidated metabolite NaAD was not detected in bacteria, as previously observed in mammalian blood following the administration of NR by Trammell and colleagues (Trammell et al. 2016a), other deamidated
NAD+ precursors such as NA, NaR and NaMN were also detected in the supernatant following
supplementation with NMN in bacteria, suggesting they may also serve as extracellular
precursors to neighbouring or host cells in the GIT.
Study Limitations
The findings from this study relied on the detection of NAD+ metabolites to infer enzymatic
routes of NMN uptake and utilisation in E. coli bacteria, specifically via dephosphorylation to
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NR as well as hydrolysis to NAM. Measuring the enzymatic activity and/or expression of these
proteins would provide important insights into the interpretation of these metabolomics data,
and future work should aim to incorporate this (Burns, Mendz, and Hazell 1998). Detection of
metabolites in this study was mainly supported and confirmed by the incorporation of labelled
ribose and nicotinamide moieties from labelled NMN isotope, NMN1, which is labelled at five
carbons of the ribose (13C) and at the base nitrogen of the pyridine ring of the nicotinamide
moiety (15C). This is an important tool which has not been used to study the bacterial
metabolism of exogenously NMN, which was an advantage of this study.
Other than the possibility that periplasmic metabolites contaminated the supernatant
during sample processing, detection of NR in the supernatant may also suggest that dephosphorylation could be occurring outside the cell. The only known 5’-nucleotidase in
bacteria, UshA, however, is located in the periplasmic compartment, and not on the outer
membrane of E. coli. An alternative explanation for these data could be that NMN undergoes modification as a result of constituents present in growth broth, rather than through the action of bacterial enzymes. Future studies should include measurements for the stability of labelled
NMN in cell-free broth to exclude this possibility. There is some precedence for the concept that culture media could degrade or alter NMN. A recent study (Kulikova et al. 2019) measured
the impact of culture conditions on the chemical and enzymatic stability of NAD precursors,
showing that the presence of 10% fetal bovine serum (FBS) in mammalian cell culture media
resulted in nucleotide pyrophosphatase and 5’-nucleotidase activity, which degraded several
NAD+ metabolites, including the dephosphorylation of NMN, however the presence of FBS in
mammalian tissue culture is a very different scenario to the bacterial culture described here.
While not investigated in this study, it is a possibility, that NMN is deamidated and/or
dephosphorylated through non-enzymatic means including by chemical degradation from
compounds present in the growth media. The culture medium, Luria-Bertani (LB) broth used
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to grow E. coli cells in this study was prepared with standard ingredients; 10 g/L tryptone, 5 g/L yeast extract and 10 g/L sodium chloride in deionized water. While it is possible that enzymes present in yeast extract could metabolise NMN, this broth undergoes autoclaving prior to use, and this heating would be expected to degrade any NAD consuming enzymes.
Regardless, future studies should incorporate a cell-free control group supplemented with
NMN to confirm this.
Another limitation of this study is the use of a single strain of a single bacterial species, in this case the OP50 strain of E. coli. This strain was chosen due to its immediate availability and ease of use. This species encodes enzymes involved in the de novo (NadD and NadE), salvage (NadR) and deamidation (PncB and PncC) pathways of NAD+ biosynthesis , and in
this study was considered to be a generic model organism for bacterial NAD metabolism. In
contrast, in the mammalian gut microbiome there are over 1000 species of bacteria and E. coli has been reported to account for only a small proportion (0.1%-2.1%) of the microbial population in the GIT, which is mainly dominated by Bacteroidetes and Firmicutes, which
together account for over 50-60% of the gut microbiome (D’Argenio and Salvatore 2015;
Huttenhower et al. 2012; Lynch and Pedersen 2016; Segata et al. 2012). Exploring the
differences in NMN uptake and utilisation pathways between different bacterial strains may
elucidate further interactions between cells and between species, which will be important to
understand in the context of stark changes in bacterial diversity based on various health and
disease conditions, including ageing (Biagi et al. 2010; D’Argenio and Salvatore 2015; Le
Chatelier et al. 2013; Ley et al. 2006; Mariat et al. 2009; Qin et al. 2012; Tilg and Kaser 2011;
Yatsunenko et al. 2012). These changes could in turn influence the bioavailability of NAD+
precursors to the host.
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Host-microbiome interaction in NMN metabolism
5.1 Introduction
Nicotinamide adenine dinucleotide (NAD+) is an essential redox cofactor central to processes
involved in energy metabolism such as glycolysis, the TCA cycle and fatty acid oxidation
(Braidy et al. 2018; Kulkarni and Brookes 2019). It is also a key substrate consumed by NAD+-
dependent enzymes such as the sirtuins (Imai et al. 2000b) and poly(ADP-ribose) polymerases
(PARPs) (Chambon, Weill, and Mandel 1963) which are important epigenetic regulators of
gene expression and genome stability (Haigis and Sinclair 2010) and DNA repair (de Murcia
and de Murcia 1994). Given the essential role of this metabolite to cell survival, the decline in
NAD+ that occurs during biological ageing (Braidy et al. 2011a; Camacho-Pereira et al. 2016;
Clement et al. 2019; Gomes et al. 2013; Massudi et al. 2012b) and in various disease states
(Frederick et al. 2016; Zhou et al. 2016) is considered to be a potential cause of declining health. Boosting levels of NAD+ through pharmacological supplementation with NAD+
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Chapter 5 Host-microbiome interactions in NMN metabolism precursors has gained attention as a promising strategy for therapeutic intervention (Aman et
al. 2018; Connell, Houtkooper, and Schrauwen 2019; Madeo et al. 2019; Mouchiroud,
Houtkooper, and Auwerx 2013; Rajman, Chwalek, and Sinclair 2018; Sultani et al. 2017;
Yoshino, Baur, and Imai 2018).
Boosting NAD+ levels through supplementation with NAD+ precursors such as nicotinamide mononucleotide (NMN) and nicotinamide riboside (NR) are emerging as promising strategies to combat various pathological conditions (Belenky et al. 2007; Cantó et al. 2012; Caton et al. 2011; Chi and Sauve 2013; Das et al. 2018; de Picciotto et al. 2016;
Diguet et al. 2018; Gong et al. 2013; Johnson, Wozniak, and Imai 2018; Lu et al. 2014a; Mills et al. 2016; Park et al. 2016; Pham et al. 2019; Tarantini et al. 2019; Trammell et al. 2016c;
Uddin et al. 2017; Wang et al. 2016b; Wei et al. 2017; Yamamoto et al. 2014b; Zhang et al.
2017). In comparison to other well-known NAD+ precursors such as nicotinic acid (NA) and
nicotinamide (NAM), collectively termed niacin, the use of NMN and NR has several
advantages. Unlike NA, the use of NMN and NR does not lead to uncomfortable flushing in
patients which is caused by the activation of the G-protein-coupled receptor, GPR109A (Benyó et al. 2005; Tunaru et al. 2003). At high doses, NAM can inhibit the sirtuins, a family of NAD+-
dependent histone and protein deacetylase enzymes involved in the regulation of gene
expression through silencing (Avalos, Bever, and Wolberger 2005; Bitterman et al. 2002), though NMN and NR do not display this effect.
Historically, niacin was used to treat pellagra, a disease prevalent in malnourished populations, caused by a chronic deficiency in dietary vitamin B3 and/or tryptophan, which are precursors for NAD+ synthesis (Alport, Ghalioungui, and Hanna 1938; Fouts et al. 1937;
Goldsmith et al. 1952; Spies, Cooper, and Blankenhorn 1938). NA has also been used to treat individuals with dyslipidaemia and hypercholesterolemia (Achor et al. 1958; Altschul, Hoffer, and Stephen 1955) but high doses lead to painful flushing side effects, which in some cases
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Chapter 5 Host-microbiome interactions in NMN metabolism manifest as a deterioration in glycaemic and plasma uric acid control, leading to hyperglycaemia and hyperuricaemia, further limiting its therapeutic application (Benyó et al.
2005; Garg and Grundy 1990). The enzymatic synthesis of NAD+ from NAM involves the addition of ribose-5-phosphate from phosphoribose pyrophosphatase by the enzyme nicotinamide phosphoribosyltransferase (NAMPT) to form NMN, in a reaction which is considered the rate-limiting step in the salvage pathway (Revollo, Grimm, and Imai 2004). The enzyme nicotinamide mononucleotide adenylyltransferase (NMNAT1-3) converts NMN into
NAD+ in a reaction that uses ATP as a donor for AMP, yielding free pyrophosphate (Berger et al. 2005; Garavaglia et al. 2002; Schweiger et al. 2001; Zhang et al. 2003), while NAD+ synthesis from NR involves its phosphorylation into NMN via nicotinamide riboside kinase
(NRK1/2) (Tempel et al. 2007) prior to acting as a subsrate for NMNAT1-3. As such, NMN and NR are ideal candidates for raising NAD+ levels in a manner which is, to our knowledge, relatively safe and absent of any harmful and or/undesirable side effects, bypassing the rate- limiting step of NAD+ biosynthesis in the salvage pathway (Airhart et al. 2017b; Conze,
Brenner, and Kruger 2019; Dellinger et al. 2017; Irie et al. 2019; Martens et al. 2018; Mills et al. 2016).
When considering the translatability of NAD+ precursors in a clinical setting, it is important to understand of mechanisms of uptake and utilisation in the cell. The route of administration significantly impacts these mechanisms and influences the efficacy of treatment. A recent study by Liu and colleagues compared the metabolic fate of NMN when delivered intravenously versus orally, and found only intravenously delivered NMN remained intact in tissues, while the oral route resulted in breakdown into NAM in the liver (Liu et al.
2018). For orally delivered compounds to have an effective systemic effect, they must survival the gastrointestinal environment, where bacterial populations thrive and overcome the ‘first pass effect’, which refers to the reduction in drug concentration at the target site due to initial
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Chapter 5 Host-microbiome interactions in NMN metabolism metabolism at a specific location in the body (Herman and Santos 2019). The first pass effect is often associated with the liver, a major site of drug metabolising enzymes such as cytochrome
P450 (CYP450) (Anzenbacher and Anzenbacherova 2001), but the microbiome in the gastrointestinal environment is also a major contributor to xenobiotic metabolism (Clarke et al.
2019; Collins and Patterson 2019; Spanogiannopoulos et al. 2016; Wilson and Nicholson
2017). Whether the microbiome plays a role in the metabolism of NAD+ precursors is not well understood, and may be important for future therapeutic applications.
The requirement for NAD+ as a co-factor is conserved across evolution, however the steps to NAD biosynthesis vary between kingdoms. In bacteria, NAD+ synthesis includes enzymatic steps that are, to our current knowledge, absent in mammalian NAD+ metabolism.
Bacteria use the NMN deamidase PncC to catalyse the conversion of NMN to its acidic form, nicotinic acid mononucleotide (NaMN) (Galeazzi et al. 2011; Sorci et al. 2014), a step which is not present in mammals. Bacteria then convert NaMN to nicotinic acid adenine dinucleotide
(NaAD) through the NaMN adenylyltransferase NadD (Mehl, Kinsland, and Begley 2000;
Preiss and Handler 1958b), which is then amidated via the NAD synthetase, NadE (Spencer and Preiss 1967; Willison and Tissot 1994), utilising ammonia as a nitrogen source for the final step in NAD+ synthesis, instead of glutamine as in mammals. Interestingly, a recent study by
Trammell and colleagues revealed the striking appearance of NaAD following oral administration of NR, the direct precursor to NMN, in mice and in a human volunteer
(Trammell et al. 2016a). Given that canonical pathways of NAD+ biosynthesis in mammals do not have a step that would allow NR to feed into the synthesis of NaAD, this finding is surprising. One option would be the existence of a mammalian NMN deamidase, however this has been controversial (Magni et al. 2008; Rongvaux et al. 2003; Sarma, Rajalakshmi, and
Sarma 1961). An alternative explanation is that the appearance of NaAD from NR could potentially occur through engaging bacterial deamidase enzymes such as PncC in the gut. NR
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Chapter 5 Host-microbiome interactions in NMN metabolism can be phosphorylated to NMN via NR kinase enzymes (NRK1/2) (Tempel et al. 2007), and
one hypothesis is that NMN could then undergo deamidation to NaMN by the bacterial enzyme
PncC through interactions with the gut microbiome. These bacterial enzymes could facilitate
host NAD+ metabolism, or conversely, limit the availability of NAD+ precursors to the host through direct consumption. Given the highly conserved requirement for NAD+ in metabolism across evolution (Gazzaniga et al. 2009), it is conceivable that microbial species in the gut may consume and/or metabolise NAD+ precursors obtained from dietary sources, or when provided by exogenous supplements. Another alternative explanation could be that NAD+ undergoes
deamidation to NaAD, however the amidation of NaAD into NAD+ by NAD synthetase is, to
our knowledge, an irreversible reaction and for this to proceed in the opposite direction would
be thermodynamically unfavourable. We therefore propose the deamidation of NMN as a
reasonable hypothesis to explain the initial observation by Trammell and colleagues and
further, examine the role of the microbiome in NMN metabolism.
In addition to the role of the microbiome in NAD+ precursor metabolism, another key
question in the field is the mechanism of NMN transport into cells. There are two proposed
mechanisms, which will be referred to in this study as ‘direct’ or ‘indirect’ mechanisms of
NMN transport (Figure 5.12). The indirect mechanism involves the extracellular
dephosphorylation of NMN to NR by the ecto-5’-nucleotidase, cluster of differentiation 73
(CD73) (Grozio et al. 2013). Following dephosphorylation of NMN, NR enters the cell via
SLC29A1 and SLC29A2, which are the primary members of a family of equilibrative
nucleoside transporters (ENT1-3) selective for purine and pyrimidine nucleosides, some
nucleobases and adenosine (Baldwin et al. 2004; Young et al. 2008). Once transported into the
cell, NR kinases (NRK1/2) phosphorylate NR into NMN (Bieganowski and Brenner 2004;
Ratajczak et al. 2016), which is used by NMNAT enzymes as a substrate for NAD+ synthesis
(Berger et al. 2005; Garavaglia et al. 2002; Schweiger et al. 2001; Zhang et al. 2002). In
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Chapter 5 Host-microbiome interactions in NMN metabolism contrast, a competing theory suggests that NMN undergoes direct, intact transport via a
dedicated NMN transporter which is highly expressed in the gut epithelium (Grozio et al.
2019a) where NAD+ is synthesised in one enzymatic step by the enzymes NMNAT1-3. In addition to addressing our hypothesis of NMN deamidation via bacterial enzymes in the gut, this study allowed us to also gain insights into which of these two mechanisms is more dominant, particularly in intestinal tissue where the putative NMN transporter is highly expressed (Grozio et al. 2019a). To address this, we paired the oral administration of strategically labelled NMN isotopes to ablation of the gut microbiome through antibiotic treatment to investigate the role of the microbiome in NMN metabolism. By tracing the incorporation of labelled nicotinamide and ribose moieties of NMN, we were able to investigate our hypotheses surrounding the contribution of the gut microbiome to the metabolism of orally administered NMN compounds and the mechanism of transport of NMN into cells. In these investigations, our emphasis was on the intestinal tissue, as our questions pertained to the contributions of the microbiome and the role of NMN transporters in the gut.
5.1.1 Rationale for NMN isotope design
In order to assess whether the gut microbiome contributes to the metabolism of orally delivered
NMN, isotope tracing was used to examine its routes of assimilation and in particular, to differentiate exogenously supplemented material from endogenous species of NAD+
metabolites that were already present in each sample. A key aspect was the strategic design of
NMN isotopes that would answer specific questions regarding the deamidation and
assimilation of NMN into the cell. To investigate this, two separate NMN isotopes were
designed that would provide complementary evidence for our hypotheses.
The first of these isotopes, designated as NMN1, was labelled with 13C-carbon at all
five carbon positions of the ribose, as well as with 15N-labelled at the pyridinyl ring of the
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Chapter 5 Host-microbiome interactions in NMN metabolism nicotinamide moiety, for an overall mass shift of M+6 (Figure 5.1 A). This isotope was co-
administered with 15N-amide-labelled glutamine (Figure 5.1 B).
Figure 5.1 Labelled isotope compounds 13 15 (A) M+6 labelled NMN1; labelled at ribose ( C5) and base ( N) of pyridine ring position. (B) M+1 labelled glutamine; labelled at amide (15N) position. 13 15 15 (C) M+7 labelled NMN2; labelled at ribose ( C5), base ( N) and amide ( N) position.
The rationale for this experiment was that if NMN undergoes deamidation prior to its
incorporation into NAD+, it would be metabolised via the de novo pathway, where the last step
involves the amidation of NaAD into NAD+ by the enzyme NAD synthetase (NADSYN). In
mammals, NADSYN catalyses the transfer of an amine group (NH2) from glutamine to NaAD,
+ replacing the hydroxyl group (R-OH) with an amide (R-NH2), to form NAD (Figure 5.2). By
generating an isotope labelled NMN with an M+6 mass shift and administering in the presence
of 15N-amide-labelled glutamine, the detection of NAD+ with an overall mass shift of M+7, at
quantities that exceed the expected background for naturally occurring 13C, would provide
evidence for the metabolism of NMN via the de novo route. To determine whether microbial
metabolism would contribute to this proposed step, the isotope was delivered to mice treated
with antibiotics to deplete their intestinal microbiome. If this proposed deamidation step was
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Chapter 5 Host-microbiome interactions in NMN metabolism mediated by the microbiome, it could be expected that there would be a reduction in the
appearance of M+7 labelled NAD+, while if this step was mediated by an unknown mammalian
host enzyme, there would be no reduction in M+7 labelling of NAD+.
The second of these isotopes, designated as NMN2, was labelled at all five carbons of the ribose moiety with 13C with a mass shift of M+5, and was also labelled at the pyridine ring
and amide with 15N on the nicotinamide moiety for an additional M+2 mass shift, giving it a final mass shift of M+7 (Figure 5.1 C). To answer our hypothesis, if NMN underwent deamidation prior to its incorporation into NAD+, we would anticipate the loss of one mass
unit, as the 15N label at the amide group of the nicotinyl moiety would be displaced by a
hydroxyl (-OH) group forming M+6 labelled NaMN as in bacteria, and NAD+ would carry an overall mass shift of M+6, rather than M+7 (Figure 5.3). NMN2 was also used to explore other aspects of NMN metabolism, such as NMN transport through direct and/or indirect mechanisms, by tracing its appearance into several metabolic fates as illustrated in Figure 5.4.
Given that one mechanism for NMN transport involves its extracellular dephosphorylation by
CD73 prior to uptake, measuring the appearance of M+7 labelled NR would quantify the relative contribution of this mechanism to NMN uptake and NR availability.
As illustrated in Figure 5.4, we proposed that NMN2 would result in the formation of
M+2 labelled free nicotinamide (NAM), due to the cleavage of M+7 labelled NAD+ by NAD+
consuming enzymes such as the PARPs and sirtuins, however, we do not exclude the possibility
that M+2 labelled NAM could represent cleavage of NMN at its labile glycosidic bond. This
M+2 labelled NAM could then be re-incorporated into NAD+ via the recycling pathway and
the actions of the enzymes NAMPT and NMNAT1-3, resulting in the appearance of M+2
labelled NAD+. NMN2 further allowed us to explore the hypothetical involvement of bacterial
deamidases, PncA and PncC, in the NAD+ metabolome. PncA converts nicotinamide (NAM)
to nicotinic acid (NA) (Frothingham et al. 1996) while PncC uses NMN as a substrate to form
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Chapter 5 Host-microbiome interactions in NMN metabolism NaMN (Galeazzi et al. 2011). Deamidation of NMN2 via PncC would form M+6 labelled
NaMN and in the presence of NMNAT1-3 and glutamine-dependent NADSYN, result in M+6
labelled NaAD and NAD+, respectively. Cleavage of M+6 labelled NAD+ by NAD+-
consuming enzymes would result in M+1 labelled NAM, which has two different fates. The
first involves normal recycling through the salvage pathway, where M+1 labelled NMN and
NAD+ are formed via NAMPT and NMNAT1-3 enzymes. The second involves deamidation
via PncA, forming M+1 labelled NA which may be used to synthesise M+1 labelled NaMN,
NaAD and NAD+ as part of the Preiss-Handler pathway (Figure 5.4). If bacterial enzymes meaningfully contribute to the NAD+ metabolome, we expect antibiotic treatment to reduce
M+6 labelling of NaMN as well as reduce M+1 labelling of NA, following the loss of 15N- labelled amide from NMN2. If this were the case, given that NA is a precursor to NaMN in the
Preiss-Handler pathway, M+1 labelled NA could result in M+1 labelling of NaMN, NaAD and
NAD+.
The NMN2 isotope was better suited to addressing these metabolic fates than the
NMN1 isotope, as an M+1 mass shift in free nicotinamide from the NMN1 isotope could be
difficult to distinguish from background 13C isotopomers of nicotinamide at low levels of
incorporation.
As one of the main aims of this study was to investigate whether the microbiome was
contributing to the metabolism of NMN, findings from these experiments are shown
predominantly in the intestinal tissue. The intestinal tissue samples were derived from the entire
length of the gastrointestinal tract including the small and large intestine as well as the proximal
and distal colonic segments (excluding cecum).
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Figure 5.2 Schematic of the NMN deamidation and re-amidation hypothesis using NMN1 and 15N-glutamine. Incorporation of isotope labels into M+6 labelled NaMN, NaAD and M+7 labelled NAD+, following administration with NMN1 (50 mg/kg, oral gavage) and 15N-glutamine (5.0 mmoles/kg, approximately 735.7 mg/kg, intraperitoneal injection).
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Chapter 5 Host-microbiome interactions in NMN metabolism
Figure 5.3 Schematic of NMN deamidation hypothesis using NMN2. Loss of 15N-amide and incorporation of isotope labels into M+6 labelled NaMN, NaAD and NAD+, following administration with NMN2 (50 mg/kg, gavage).
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Chapter 5 Host-microbiome interactions in NMN metabolism
Figure 5.4 Schematic for the potential biosynthesis pathways and NAD+ metabolites involved in the utilisation of NMN2. NMN2 (box on left) was labelled with 13C at all 5 carbon positions of the ribose moiety (highlighted in red) and with 15N at the base (highlighted in blue) and amide (highlighted in orange) of the nicotinamide moiety for a total mass shift of M+7. The expected mass shift for each species depending on the potential route of utilisation is shown, with steps catalysed by mammalian enzymes represented by solid black lines, and steps that are only known to be carried out by bacterial enzymes represented by the dashed lines.
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Chapter 5 Host-microbiome interactions in NMN metabolism 5.2 Methods
5.2.1 Animal housing
All animal experiments were performed according to procedures and guidelines approved by
UNSW Animal Care and Ethics Committee (ACEC) under the ethics protocol 18/133A. The
UNSW ACEC operates under the Animal Ethics Guidelines from the National Health and
Medical Research Council (NHMRC) of Australia. All mice were housed in individually ventilated cages of no more than five mice per cage and fed a standard chow diet and drinking water ad libitum. The cages were maintained under a 12-hr light/12-hr dark cycle in a temperature-controlled room (22 ± 1℃) at 80% humidity. All cages were changed twice per week and monitored daily.
5.2.2 Antibiotic and NMN treatment regimen
Four-week-old female C57BL/6J mice were acclimatised for one week prior to treatment. At five weeks, all mice were body-weight-matched and randomly assigned into one of the following treatment groups.
Antibiotic treatment
For antibiotic treatment, mice were administered a cocktail of antibiotics consisting of vancomycin (0.5 g/L; Sigma SBR00001), neomycin (1 g/L; Sigma N6386), ampicillin (1 g/L;
Sigma A9393) and metronidazole (1 g/L; Sigma M3761) (herein, referred to as VNAM). This antibiotic cocktail was administered in the drinking water of mice for four days with the addition of sucrose (3 g/L; Bundaberg Sugar) in all treatment groups including controls to increase palatability and prevent aversion to the drinking water during antibiotic treatment
(Jimeno, Brailey, and Barral 2018). The body weight of mice was recorded during the antibiotic treatment period however, there was a reduction in water consumption in the VNAM cocktail group (Figure 5.5 A) which can increase their risk of illness or death due to dehydration
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Chapter 5 Host-microbiome interactions in NMN metabolism (Reikvam et al. 2011). As a result, VNAM was replaced by ampicillin treatment only, with
sucrose, for an additional week which was previously shown to effectively reduce gut bacterial
density by 103-fold (Jimeno, Brailey, and Barral 2018; Ubeda et al. 2010).
NMN2 cohort
After the antibiotic treatment period, mice were orally administered an equivalent bolus of
water or NMN2 by gavage according to the following treatment groups: 1) Vehicle control
(water) (n=3), 2) Antibiotics only (Abx) (n=4), 3) NMN2 only (n=4) and 4) NMN2+Abx (n=4).
NMN1+15N-amide-glutamine cohort
After the antibiotic treatment period, mice were orally administered an equivalent bolus of
water or NMN1 by gavage and/or an intraperitoneal injection of water or 15N-amide-glutamine
according to the following treatment groups: 1) Vehicle control (water) (n=4), 2) Antibiotics
only (Abx) (n=4), 3) 15N-amide-glutamine only (n=3), 4) 15N-amide-glutamine+Abx (n=4), 5)
NMN1+15N-amide-glutamine (n=4) and 6) NMN1+15N-amide-glutamine+Abx (n=4).
The synthesis of labelled compounds, NMN1, NMN2 and 15N-amide-glutamine is detailed in
Chapter 2, Section 2.2.1. The final concentration of NMN isotopes and 15N-amide-glutamine were 50 mg/kg and 5.0 mmoles/kg (approximately 735.7 mg/kg), respectively, based on previous studies (Liu et al. 2018; Wang et al. 2019). All blood and tissue samples were collected four hours after gavage. The oral gavage was staggered by five mins between each mouse with the treatment groups alternating to avoid experimental bias during the procedure.
5.2.3 Plasma collection and metabolite extraction
For blood collection, all mice were placed under general anaesthesia with isoflurane gas maintained at 1.5-2% v/v and O2 flow set at 1-1.5 litres per minute. Blood was collected by
cardiac puncture via insertion of a 25-gauge needle through the diaphragm with at least 600 µL
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Chapter 5 Host-microbiome interactions in NMN metabolism of blood drawn from each mouse. Caution was taken to draw the blood slowly into a 1mL
syringe to prevent the collapse of heart chambers due to pressure changes. The volume of blood
obtained was immediately expelled into a 1.5 mL tube pre-filled with 10 µL of 0.5 M
ethylenediaminetetraacetic acid (EDTA) solution which was thoroughly mixed up and down
with a pipette to prevent the blood from clotting. Plasma was separated from whole blood
contents via centrifugation at 2000 g for 15 minutes at 4℃ before being transferred to a clean
pre-labelled tube and snap frozen immediately in liquid nitrogen. Samples were stored at -80℃
until further processing. On the day of sample acquisition, the following protocol was used for
extraction of NAD+ metabolites from plasma samples; 20 µL of plasma was combined with 80
µL of cold (-30℃) extraction buffer consisting of a 1:1 ratio of LC-MS-grade acetonitrile and
methanol (ACN:MeOH) containing a mixture of internal standards, methionine sulfonate
(MES), camphorsulfonic acid (CSA) and thymine-d4, as in Chapter 2, Section 2.2.1. Samples
were vortexed and centrifuged at 16,000 g for 10 minutes at 4℃ after which the supernatant
was transferred to a clean pre-labelled tube and dried under a Savant SpeedVac™ system
(Thermo Scientific) under vacuum for approximately 2 hours, as in Chapter 2, Section 2.2.3.
The resulting dry pellet was resuspended in 30 µL of LC-MS-grade water and centrifuged again as described above. The supernatant was then transferred to pre-labelled mass spectrometry vials and analysed promptly by LC-MS, as described in Chapter 2, Section 2.3. Data was extracted using in-house MATLAB scripts as described in Chapter 2, Section 2.4. For data analysis, samples were normalised by dividing the raw peak area of the sample by the raw peak area of the internal standard, thymine-d4, for each metabolite. The standard curve for each metabolite was then used to calculate the molar amount (µM) of metabolite in each sample. All graphs were generated and statistically analysed using GraphPad Prism software as described in Chapter 2, Section 2.5.
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Chapter 5 Host-microbiome interactions in NMN metabolism 5.2.4 Tissue collection, metabolite extraction and data analysis
For collection and processing of mouse tissue samples, the following protocols were used.
Livers were resected and weighed immediately before rinsing in ice-cold 1X phosphate
buffered saline (PBS) to remove any residual blood from the tissue. Intestinal tissue was
resected from the small intestine to the colon (excluding cecum) and the inside was flushed
with ice-cold 1X PBS to remove any faecal contents remaining in the intestinal tract. Both
tissues were snap frozen immediately in liquid nitrogen and stored at -80℃ until further
processing. Faecal pellets were collected from the colonic region of the intestinal tract,
immediately snap frozen in liquid nitrogen and stored at -80℃ until ready for further
processing. One day prior to sample acquisition, the liver and intestinal tissues were crushed
with a mortar and pestle on liquid nitrogen to pulverise the frozen tissues. Approximately 50
mg of liver and intestinal tissue was weighed into pre-labelled tubes containing 1.4 mm ceramic
beads (Precellys® CK14, Bertin Technologies). NAD+ metabolites were extracted from
samples as described in Chapter 2, Section 2.2.2. Briefly, 500 µL of cold (-30℃) extraction buffer (HPLC-grade acetonitrile-methanol-water mixture, 2:2:1) was added to each sample with the internal standard mixture (Chapter 2, Section 2.2.1). The samples were then homogenised using a mechanical homogeniser (Precellys® 24, Bertin Technologies) at 5500 rpm for 15 seconds at room temperature. All samples were centrifuged at 16,000 g for 10 minutes at 4℃ and the supernatant was transferred to a clean pre-labelled tube before snap freezing in liquid nitrogen and storage at -80℃. All sample extracts were dried using the speed vac system (Savant SpeedVac SPD140DDA Vacuum Concentrator, Thermo Fisher Scientific) and the dried pellet was resuspended in 50 µL of LC-MS-grade water and centrifuged once more, as described above. The supernatant was transferred to pre-labelled mass spectrometry vials and analysed promptly by LC-MS/MS as described in Chapter 2, Section 2.3. The data collected from the LC-MS/MS acquisitions were extracted and analysed as described in
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Chapter 5 Host-microbiome interactions in NMN metabolism Chapter 2, Section 2.4. For data analysis, samples were normalised by dividing the raw peak area of the sample by the raw peak area of the internal standard, thymine-d4, for each metabolite. The standard curve for each metabolite was then used to calculate the molar amount
(pmol) of metabolite in each sample and normalised again to the frozen tissue weight (mg).
Final measurements are presented as pmol/mg tissue. The data were statistically analysed using
GraphPad Prism software as described in Chapter 2, Section 2.5.
5.2.5 DNA extraction from faecal pellets
Solid faecal pellets taken from the colonic and rectal region of the gastrointestinal tract were stored in -80℃ until further processing. DNA was extracted from frozen faecal pellets using the QIAamp® PowerFecal® DNA kit (Qiagen, Cat. No. 12830-50) according to the manufacturer’s protocol. DNA concentration was determined using the NanoDrop™
(DeNovix®, DS-11 FX) and the purity of double-stranded DNA (dsDNA) was also determined by measuring the 260/280 ratio. All DNA extracts were stored at -80℃ until further processing.
5.2.6 Nanopore 16S Sequencing
Full length 16S rRNA genes were amplified by PCR using the Oxford Nanopore 16S
Barcoding Kit (SQK-RAB204; Oxford Nanopore Technologies, Oxford, UK). Briefly, 10 ng genomic DNA, 1 µL 16S Barcode (10 µM) and 25 µL LongAmp Taq 2X Master Mix (New
England Biolabs, Ipswich, MA, USA) were combined in a 50 µL reaction for PCR on a Bio-
Rad T100TM Thermal Cycler (Bio-Rad Laboratories Pty Ltd, Hercules, CA, USA). PCR cycling condition were as follows; initial denaturation at 95 ℃ for 1 minute, 25 cycles of denaturation at 95 ℃ for 20 seconds, annealing at 55 ℃ for 30 seconds and extension at 65 ℃ for 2 minutes before a final extension at 65 ℃ for 5 minutes.
PCR products were purified as per Oxford Nanopore Technologies (ONT) protocol using AMPure XP magnetic beads (Beckman Coulter, Indianapolis, IN) and DNA quantified
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Chapter 5 Host-microbiome interactions in NMN metabolism using the NanoDrop™ (DeNovix®, DS-11 FX). Barcodes were pooled to a total of 100 fmol in 10 μL of 10 mM Tris-HCl, pH 8.0 with 50 mM NaCl for library loading. Sequencing was performed using R9.4.1 ONT Flow Cells on the MinIONTM sequencing platform and data acquired using MinKNOW software version 19.10.1 (Oxford Nanopore Technologies).
5.2.7 Data Analysis for 16S sequencing
Data extraction and analysis for full length 16S sequences obtained from nanopore sequencing was kindly carried out by Timothy Chalmers. Sequencing reads acquired from the MinION runs (i.e. FAST5 data) were basecalled to fastq files using Guppy software version 3.4.4
(Oxford Nanopore Technologies). Fastq files were demultiplexed using Porechop
(https://github.com/rrwick/Porechop) and trimmed to 1400bp with Trimmomatic version 0.39
(Bolger, Lohse, and Usadel 2014). Reads were imported to QIIME2 for dereplication and chimeric reads screened and filtered from the dataset. Operational taxonomic unit clustering was completed within QIIME2 version 2019.7.0 (Bolyen et al. 2018) at 85% similarity to account for typical sequencing errors obtained from long-read sequencing. Taxonomy was assigned to reads using a pre-trained classifier on the SILVA 132 16S rRNA representative sequences. Data was imported into R version 3.6.1 with qiime2R version 0.99.13
(https://github.com/jbisanz/qiime2R) for visualisation and alpha diversity analysis using raw and rarefied data with the phyloseq version 1.30.0 (McMurdie and Holmes 2013) package. The
R package DESeq2 version 1.28.1 (Love, Huber, and Anders 2014) was used to normalise and model-fit the read counts for differential expression testing between treatment groups.
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Chapter 5 Host-microbiome interactions in NMN metabolism 5.3 Results
5.3.1 Antibiotic ablation of the gut microbiome
To interrogate the contribution of the gut microbiome to the metabolism of orally delivered
NMN, C57BL/6J female mice were exposed to a course of antibiotics for eleven days through
delivery in drinking water (Figure 5.5 A). Water intake (Figure 5.5 A) and body weight
(Figure 5.5 B) was measured throughout the treatment period in order to monitor antibiotic
consumption and animal welfare. The antibiotic cocktail including vancomycin (0.5 g/L),
neomycin (1 g/L), ampicillin (1 g/L) and metronidazole (1 g/L) (VNAM) was administered in the drinking water of mice, but after four days was associated with a severe decline in water consumption (Figure 5.5 A), though no signs of body weight loss or dehydration were observed (Figure 5.5 B). Due to the potential risks of dehydration and illness from reduced water intake for a prolonged period, the VNAM cocktail was replaced with ampicillin treatment alone, which rapidly increased water consumption within two days.
Antibiotic treatment also led to differences in the internal organs of mice including an enlargement of the cecum (Figure 5.5 D) and a reduction in the weight of spleens (Figure 5.5
E) and livers (Figure 5.5 F), with no significant reduction in body weight compared to the
controls (Figure 5.5 G). These observations are consistent with previous studies using
antibiotic treatment (Cresci, Nagy, and Ganapathy 2013; Jimeno, Brailey, and Barral 2018;
Puhl et al. 2012; Reikvam et al. 2011; Xu et al. 2017) and are also observed in germ-free mice
(Al-Asmakh and Zadjali 2015; Niimi et al. 2019; Niimi and Takahashi 2019; Smith, McCoy,
and Macpherson 2007) indicating these effects are likely due to the under-representation of
commensal bacteria in the gut, rather than due to an adverse side effect of antibiotic treatment.
No significant difference was observed when antibiotic treated mice were administered NMN,
which is expected due to the acute treatment period of four hours. Antibiotic treatment also
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Chapter 5 Host-microbiome interactions in NMN metabolism dramatically reduced the DNA concentration of mouse stool samples compared to controls
(Figure 5.5 C), inferring an overall reduction in the gut microbiome from antibiotic treatment.
To test the effects of antibiotic treatment on the make-up of the remaining species in
the gut microbiome, gut contents were subjected to microbiome sequencing. Using generic
primers, full-length 16S rRNA was amplified from samples that were normalised to the amount
of starting DNA, and subjected to long-read sequencing using an Oxford Nanopore instrument
(Benítez-Páez, Portune, and Sanz 2016; Kilianski et al. 2015; Shin et al. 2016). This yielded a
significant reduction in bacterial species diversity in stools of mice treated with antibiotics
compared to controls, with the only surviving species in antibiotic-treated mice being from the
Burkholderia genus (Figure 5.6 A). This change is highlighted by alpha diversity analysis
using the Shannon diversity index (Allen, Kon, and Bar-Yam 2009) which showed
significantly reduced diversity in antibiotic treated mice, represented here at the genus level
(Figure 5.6 B). There was no change in microbial diversity when antibiotic-treated mice were
co-administered NMN, which was expected due to the acute treatment period of 4 hours
following a single oral gavage. A similar effect was observed in antibiotic-treated mice co-
administered NMN1 and 15N-glutamine (Figure 5.7) where microbial diversity was
significantly reduced, with the Burkholderia genus remaining. Differential expression analysis
showed antibiotic treated mice had significant downregulation of genes compared to vehicle
control, though there were very little significant changes observed in NMN treated mice
compared to vehicle control (Appendix Figure A.1 and Figure A.2).
The Burkholderia genus is made up of gram-negative bacteria and are common inhabitants of soil that thrive in acidic environments (Sermswan et al. 2015; Stopnisek et al.
2014). Members of Burkholderia are well known for developing resistance to antimicrobials, largely owing to low outer membrane permeability and unique lipopolysaccharide (LPS) structure which reduces the net negative charge of the cell envelope and in turn, reduces
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Chapter 5 Host-microbiome interactions in NMN metabolism permeation of cationic antimicrobials such as polymyxins (Rhodes and Schweizer 2016). In animals and humans, there are several Burkholderia species that have emerged as opportunistic pathogens which cause severe infections in cystic fibrosis and immunocompromised patients
(Carlier et al. 2014; Mann et al. 2010; Zlosnik et al. 2015). Recent evidence, however, identified new Burkholderia species with beneficial traits that promote plant growth and stress resistance (Pinedo et al. 2015; Sheibani-Tezerji et al. 2015; Su et al. 2015) with potential use in agriculture and industry. Comparative genomics studies to identify common virulence factors in Burkholderia species reveal universal virulence factors in multiple organisms and disease models is largely host-specific and remains a challenge (Angus et al. 2014; Schwager et al. 2013; Uehlinger et al. 2009). Improved methods to better assess pathogenicity of
Burkholderia strains are warranted to identify beneficial from harmful strains which in turn may be helpful in the diagnosis and treatment of diseases affected by infection by Burkholderia species. Regardless, these data suggest that antibiotic treatment was overwhelmingly effective in reducing bacterial content, with the exception of this genus.
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Figure 5.5 The effect of antibiotic treatment in mice. The (A) water intake and (B) body weight during antibiotic treatment was recorded. Antibiotic treatment significant decreased (C) DNA stool concentration and caused (D) enlargement of the cecum, decreased (E) spleen and (F) liver weight but had no effect in (G) body weight after one week of antibiotic treatment compared to controls. For statistical analysis a two-way ANOVA with Sidak’s multiple comparisons test was performed. Data represents mean ± s.d. (n=3-5) with a p-value less than 0.05 to be considered statistically significant. *p<0.05, **p<0.01, ***p<0.001, ****p<0.0001.
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Figure 5.6 Reduced diversity of the murine faecal microbiome upon antibiotic treatment in NMN2 cohort. (A) Stacked bar plots represent the total reads and relative abundance of bacterial taxa at the genus level for mice treated with vehicle control, antibiotics (Abx), NMN2, or NMN2+Abx, measured by full length 16S rRNA Nanopore sequencing. Taxa with < 0.1% abundance grouped together. (B) Shannon diversity measures of murine gut microbiome by Wilcoxon rank sum test. Data presented as mean ± s.d. (n=3-4). *p < 0.05
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Figure 5.7 Reduced diversity of the murine faecal microbiome upon antibiotic treatment in NMN1 cohort. (A) Stacked bar plots represent the total reads and relative abundance of bacterial taxa at the genus level for mice treated with vehicle control, antibiotics (Abx), 15N-glutamine, 15N- glutamine+Abx, NMN1+15N-glutamine, and NMN1+15N-glutamine+Abx, measured by full length 16S rRNA Nanopore sequencing. Taxa with < 0.1% abundance grouped together. (B) Shannon diversity measures of murine gut microbiome by Wilcoxon rank sum test. Data presented as mean ± s.d. (n=3-4). *p < 0.05
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Chapter 5 Host-microbiome interactions in NMN metabolism 5.3.2 Antibiotic treatment increases NAD+ metabolite and NMN availability in the
intestinal tissue of mice.
Due to its greater mass shift and therefore ability to discriminate between species, we initially
used the NMN2 isotope for the first part of this investigation. Following antibiotic treatment,
mice were orally administered by gavage a bolus of either water or NMN2 (50 mg/kg). Four
hours later, animals were euthanased and tissues rapidly collected and snap frozen. NAD+
metabolites were detected using the optimised LC-MS/MS method developed in Chapter 3.
In addition to NMN2, NMN1 was also administered in mice combined with 15N-amide-
glutamine (Figure 5.1 B), a necessary cofactor in the last step of the Preiss-Handler pathway
where NaAD is converted into NAD+ via the enzyme NAD synthetase (NADSYN).
Incorporation of all labels from NMN1 (M+6) and 15N-amide-glutamine (M+1) into NAD+,
can be detected by LC-MS/MS as M+7 labelled NAD+ and may provide evidence to suggest
NMN deamidation into NaMN and its re-amidation through the Preiss-Handler pathway. If this
occurs through bacterial deamidases in the gut, mice treated with antibiotics would show a
dampening of this effect.
In the intestinal tissue, an increase in the abundance of unlabelled NAD+ metabolites,
NR, NAD+, NAM and NMN, was observed in mice treated with antibiotics alone (Figure 5.7
A-D), even without the exogenous supplementation with NMN. This was a surprising observation as our original hypothesis anticipated a reduction in metabolites with antibiotic treatment, rather than an increase, if bacterial enzymes in the gut facilitated NAD+ biosynthesis.
In contrast to our hypothesis, these results suggest that bacteria in the gut may be competing
with the host for baseline uptake of trace nutrients from dietary sources (Chi and Sauve 2013;
Mielgo-Ayuso et al. 2018; Mills et al. 2016; Yao et al. 2011), implicating the involvement of
the gut microbiome in host NAD+ availability.
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Chapter 5 Host-microbiome interactions in NMN metabolism In addition to differences in the levels of NAD+ metabolites derived from dietary
sources, this “competition” relationship between the microbiome and the host was tested
through treatment with an exogenous bolus of NMN, effectively overriding the host-
microbiome competition for relatively scarce dietary NAD+ precursors. The labelled species that may be detected for each of these metabolites can be categorised into two distinct groups, which are referred to here as ‘complete’ (Figure 5.7 E-H) or ‘partial’ (Figure 5.7 I-L) labelling of metabolites species. Complete labelling of metabolites represents the incorporation of both labelled ribose and nicotinamide moieties from the NMN2 isotope, while partial labelling of metabolites represents the incorporation of labelled nicotinamide species formed from the
NAD+ degradation from NAD+-consuming activities. As modelled in Figure 5.4, the
contribution of the microbiome in the gut the specific tracing of metabolites allow the
contribution of the gut microbiome to the metabolism of exogenously supplemented NMN in
the host to be determined. The incorporation of M+7 label into the NAD+ metabolome occurred
almost exclusively in antibiotic treated animals as denoted by the red bars in Figure 5.7 E-H.
These data suggest that even at this small bolus dose, the gut microbiome is still in competition
with the host for NAD+ metabolites, limiting the availability of supplemented NMN, and/or
impeding its intestinal absorption in the host. These effects were extended beyond the
gastrointestinal tract. The simultaneous increased abundance of partially labelled M+2 NAD+
(Figure 5.7 J) and NMN (Figure 5.7 L) in intestinal tissue, M+7 NAD+ (Figure 5.8 F) and
M+2 NAM (Figure 5.8 G) in the liver and M+2 NAM in the plasma (Figure 5.9 B) add weight
to the availability of NMN to the host.
The effect of antibiotic treatment on NAD+ metabolites detected in the intestinal and
liver homogenates from the NMN2 cohort was also reflected in the cohort of mice administered
NMN1, which has an overall mass shift of M+6. Similar to observations in the NMN2 cohort,
antibiotic treatment significantly increased the endogenous unlabelled pool of NAM in
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Chapter 5 Host-microbiome interactions in NMN metabolism intestinal tissues with a similar, but non-significant trend observed in NR, NAD+ and NMN
(Figure 5.10 A-D). In the liver, unlabelled NAD+ and NAM significantly increased with
antibiotic treatment with a similar, but non-significant, trend observed in unlabelled NR and
NMN (Figure 5.11 A-D). A significant increase in the incorporation of M+6 label into NR,
NAD+ and NMN was detected in intestinal homogenates following the administration of
NMN1 to antibiotic treated mice and denoted by the red bars in Figure 5.11 E, F and H. The
turnover of M+6 labelled NAD+ in the intestine was also affected by antibiotic treatment,
forming M+1 labelled NAM at the pyridine base, which significantly increased exclusively in
antibiotic treated mice following a bolus dose of NMN1 (Figure 5.11 G). The partially labelled
metabolites, M+5 NR (Figure 5.11 I), M+1 NAM labelled at the amide from 15N-glutamine
(Figure 5.11 K) and M+1 NMN (Figure 5.11 L), also significantly increased following
antibiotic treatment, highlighting the effect of the microbiome on NMN utilisation as well as
on metabolic processes involved in NAD+ degradation and biosynthesis. Unlike the intestinal
tissue, the complete and partial incorporation of labels into NAD+ metabolites detected in the
liver following administration with NMN2 (Figure 5.8) or NMN1 (Figure 5.11), were not as
striking. This could be due to the majority of metabolites being degraded and/or consumed by
bacterial enzymes in the intestinal environment prior to reaching the liver. Alternatively,
antibiotic treatment may have induced damage to the liver, as well as the spleen (Figure 5.5
E) and possibly the integrity of the intestinal epithelium (though was not investigated here), as there was a reduction in liver weight in antibiotic treated mice compared to controls (Figure
5.5 F, above). Collectively, the results from both cohorts highlight the significant increase in the availability of orally delivered NMN and NAD+ metabolites to host tissues with antibiotic
treatment, likely mediated by changes to the microbiome. It is possible that the microbiome
acts in competition with the host for NAD+ metabolites for its own growth and proliferation,
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Chapter 5 Host-microbiome interactions in NMN metabolism and/or impeding intestinal absorption of NMN in the host, however more investigations need
to be conducted in order to further support these claims.
Metabolites detected in the blood represent those capable of circulating between tissues
such as the intestines and the liver, acting as an extracellular source of NAD+ precursors to ensure neighbouring cells and tissues can maintain intracellular NAD+ levels. The main NAD+
metabolite detected in the plasma of NMN administered mice was nicotinamide (NAM) which
can be derived from NAD+-consuming activities or from breaking the glycosidic bond between
the ribose and nicotinamide moieties of NMN2, yielding M+2 NAM (Figure 5.9 B), or NMN1,
yielding M+1 NAM (Figure 5.12 B). Treatment with antibiotics had little impact on these
labelled NAM isotopologues (red bars) in the plasma, whereas in the intestines and liver a
significant increase in M+2 NAM (Figure 5.7 G for intestines and Figure 5.8 G for liver) and
M+1 NAM (Figure 5.10 G for intestines) was observed following an exogenous bolus of
NMN2 or NMN1, respectively, indicating NAM metabolism in these tissues is significantly
impacted by ablation of the microbiome through antibiotic treatment.
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Figure 5.8 An increase in amidated NAD+ metabolites detected with antibiotic treatment in the intestinal tissue after NMN2 administration. Five-week old C57BL/6 mice were treated with or without antibiotics (Abx) before receiving a single oral gavage of vehicle (water) or NMN2 (50 mg/kg). The abundance of NAD+ metabolites in the intestinal tissue, shown above, was analysed using LC-MS/MS. (A-D) represents the abundance of unlabelled species. (E-H) represents complete labelling while (I-L) represents partial labelling of NR, NAD+, NAM and NMN. Data were analysed by two-way ANOVA with Sidak’s multiple comparisons test and represent mean ± s.d. (n=3-4 mice per group). **p<0.01, ***p<0.001, ****p<0.0001.
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Figure 5.9 Detection of amidated NAD+ metabolites with antibiotic treatment in the liver after NMN2 administration. Five-week old C57BL/6 mice were treated with or without antibiotics (Abx) before receiving a single oral gavage of vehicle (water) or NMN2 (50 mg/kg). The abundance of NAD+ metabolites in the liver, shown above, was analysed using LC-MS/MS. (A-D) represents the abundance of unlabelled species. (E-H) represents complete labelling while (I-L) represents partial labelling of NR, NAD+, NAM and NMN. Data were analysed by two-way ANOVA with Sidak’s multiple comparisons test and represent mean ± s.d. (n=3-4 mice per group). *p<0.05, **p<0.01, ***p<0.001.
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Figure 5.10 Detection of nicotinamide (NAM) with antibiotic treatment in the plasma after NMN2 administration. Five-week old C57BL/6 mice were treated with or without antibiotics (Abx) before receiving a single oral gavage of vehicle (water) or NMN2 (50 mg/kg). The abundance of NAM (nicotinamide) in the plasma, shown above, was analysed using LC-MS/MS. (A) M+0 represents unlabelled NAM which is endogenously present in the plasma, while (B) M+2 labelled NAM represents NAM labelled with 15N at the pyridine ring and amide functional group. It can be formed from the degradation of M+7 labelled NAD+, involving cleavage at the glycosidic bond between the nicotinamide and ribose moieties from NAD+-consuming activities. Data were analysed by two-way ANOVA with Sidak’s multiple comparisons test and represent mean ± s.d. (n=3-4 mice per group). *p<0.05, **p<0.01, ****p<0.0001.
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Figure 5.11 An increase in amidated NAD+ metabolites detected with antibiotic treatment in the intestinal tissue after NMN1 administration. Five-week old C57BL/6 mice were treated with or without antibiotics (Abx) before receiving a single oral gavage of vehicle (water) or NMN2 (50 mg/kg). The abundance of NAD+ metabolites in the intestinal tissue, shown above, was analysed using LC-MS/MS. (A-D) represents the abundance of unlabelled species. (E-H) represents complete labelling while (I-L) represents partial labelling of NR, NAD+, NAM and NMN. Data were analysed by two-way ANOVA with Sidak’s multiple comparisons test and represent mean ± s.d. (n=3-4 mice per group). *p<0.05, **p<0.01, ***p<0.001.
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Figure 5.12 Detection of amidated NAD+ metabolites with antibiotic treatment in the liver after NMN1 administration. Five-week old C57BL/6 mice were treated with or without antibiotics (Abx) before receiving a single oral gavage of vehicle (water) or NMN2 (50 mg/kg). The abundance of NAD+ metabolites in the liver, shown above, was analysed using LC-MS/MS. (A-D) represents the abundance of unlabelled species. (E-H) represents complete labelling while (I-L) represents partial labelling of NR, NAD+, NAM and NMN. Data were analysed by two-way ANOVA with Sidak’s multiple comparisons test and represent mean ± s.d. (n=3-4 mice per group). *p<0.05.
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Chapter 5 Host-microbiome interactions in NMN metabolism
Figure 5.13 Detection of nicotinamide (NAM) with antibiotic treatment in the plasma after NMN1 administration. Five-week old C57BL/6 mice were treated with or without antibiotics (Abx) before receiving a single oral gavage of vehicle (water) or NMN2 (50 mg/kg). The abundance of NAM in the plasma, shown above, was analysed using LC-MS/MS. (A) Unlabelled NAM increased with NMN1 administration which may be due to a frameshift increase in NAD+ metabolites following an exogenous bolus of NMN. (B) No significant change was observed in antibiotic treated mice. (C) Labelling of NAM at the amide comes from 15N-amide-labelled glutamine (15N-gln), however, this could be due to a frameshift increase in NMN rather than from 15N- gln as no change was observed between mice injected with 15N-gln (in green) to controls (in grey). Data were analysed by two-way ANOVA with Sidak’s multiple comparisons test and represent mean ± s.d. (n=3-4 mice per group). *p<0.05.
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Chapter 5 Host-microbiome interactions in NMN metabolism 5.3.3 Evidence for NMN uptake facilitated by dephosphorylation into NR
The design of our isotope labelled NMN allowed us to obtain insights into the proposed
mechanisms of NMN transport by intestinal cells in the gut. The two proposed mechanisms for
NMN uptake are either directly via a dedicated transporter (Grozio et al. 2019a) or indirectly,
via dephosphorylation into NR prior to uptake where NR is re-phosphorylated into NMN by
NR kinases (NRK1/2) (Ratajczak et al. 2016). These competing mechanisms are illustrated in
Figure 5.13. Both the NMN transporter, SLC12A8, and the ecto-5’-nucleotidase, CD73, which
dephosphorylates NMN into NR, sit on the apical side of the intestinal epithelium, and for this
reason the intestinal tissue was a key focus of this investigation. NR can enter cells via an
equilibrative nucleoside transporter (ENT), SLC29A1 or SLC29A2 (Baldwin et al. 2004;
Young et al. 2008), which is embedded in the plasma membrane. It is possible that these
mechanisms run parallel to each other, rather than being mutually exclusive events. The
question, however, is the degree to which each mechanism contributes to the availability of
intracellular NMN, or whether these are condition- or tissue-specific events. If the direct route via SLC12A8 prevailed, we would expect to see significant intact (M+7) NMN incorporation relative to the endogenous (M+0) pool, and similarly for downstream products that may include
NR.
Ratios between the ‘complete’ and unlabelled isotopologues are reported here as proxies for the relative presence and turnover of metabolites (Figure 5.14). These ratios capture
the dilution of the existing endogenous pool by the influx of supplemented NMN and NMN-
derived products. Strikingly, we observed strong labelling of the NR, but not NMN pools
(Figure 5.14 A). Isotope (M+7) labelling of the NR pool following NMN2 treatment was three-
fold greater than endogenous NR (M+0), this increased to a ten-fold enrichment in animals
treated with antibiotics. In stark contrast, only around 5% of the NMN pool was M+7 labelled,
representing a small proportion of labelled NMN detected intact. The inability of exogenous
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Chapter 5 Host-microbiome interactions in NMN metabolism NMN to displace the endogenous NMN pool, combined with the surge of NR derived directly from supplemented NMN, provides strong evidence that the availability of intracellular NMN bypasses direct transport. These data would instead support the dephosphorylation of NMN into NR to facilitate its intestinal absorption, as described in Figure 5.13. If direct transport of
NMN does occur, we propose it is in competition with the microbiome, as the M+7 labelling that was detected only occurred in the intestinal tissue of antibiotic treated animals (Figure 5.7
H and Figure 5.10 H). NR also had, by far, the highest labelled-to-unlabelled ratio compared to all other metabolites (Figure 5.14 A-B). As such, these data support the uptake of NMN in the gastrointestinal tract via an NR intermediate. There are, however, important caveats to the interpretation of these data which will be discussed in detail in the discussion.
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Figure 5.14 Schematic for the direct or indirect transport of NMN in intestinal cells. Direct transport of NMN into cells may occur via the membrane bound transporter, Solute Carrier Family 12 Member 8, SLC12A8 (Grozio et al. 2019a). Intracellular NMN is then able to synthesise NAD+ via NMN adenylyltransferases (NMNAT1-3) (Berger et al. 2005). The indirect transport of NMN requires dephosphorylation of NMN into nicotinamide riboside (NR) via the ecto-5’-nucleotidase, cluster of differentiation 73 (CD73) (Grozio et al. 2013), lying on the apical side of the membrane and faces the lumen. NR may enter the cell via equilibrative nucleoside transporters (ENT1-3), SLC29A1 or SLC29A2 (Baldwin et al. 2004; Young et al. 2008). NR is then re-phosphorylated into NMN via NR kinases (NRK1/2) in the NRK pathway (Ratajczak et al. 2016) before NAD+ synthesis via NMNAT1-3 (Berger et al. 2005). NAD+-consuming enzymes such as the sirtuins and poly(ADP-ribose)-polymerases (PARPs) degrade NAD+ into NAM which can be recycled through the salvage pathway to maintain intracellular NAD+ levels. NAM is an uncharged compound small enough to diffuse across the basolateral membrane and enter the blood. It is possible for NR to also enter the blood as the NR transporters, ENT1-3, are also embedded on the basolateral membrane of mammalian cells (Baldwin et al. 2004; Young et al. 2008).
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Figure 5.15 The ratio of the labelled-to-unlabelled isotopologues of NAD+ metabolites (A) The ratio of complete labelled-to-unlabelled species in the intestinal homogenates of antibiotic treated-mice following oral administration with M+7 labelled NMN2 is shown above. The enrichment of the label into NR but not NMN is observed which increased in antibiotic treated mice. Similarly, the ratio of partially labelled-to-unlabelled species in the intestinal homogenates is shown in (B) showing enrichment of the label into NR, which is significantly greater compared to all other metabolites, including NMN. Data were analysed by two-way ANOVA with Sidak’s multiple comparisons test and represent mean ± s.d. with the mean value for each treatment group annotated above each bar (n=3-4 mice per group). *p<0.05, **p<0.01, ***p<0.001, ****p<0.0001, ns=not significant.
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Chapter 5 Host-microbiome interactions in NMN metabolism 5.3.4 NMN deamidation is independent of the microbiome.
In the canonical pathways of NAD+ biosynthesis in mammals, the amidated metabolites NMN,
NR and NAM are not converted into their deamidated counterparts nicotinic acid
mononucleotide (NaMN), nicotinic acid riboside (NaR), and nicotinic acid (NA), respectively
(refer to schematic in Figure 5.4). These deamidated forms are confined to the Preiss-Handler pathway (Preiss and Handler 1958a), the final step of which involves the enzyme, NAD synthetase (NADSYN), amidating NaAD into NAD+ (Figure 5.4). Surprisingly, other research
groups have reported that treatment with the amidated precursor, NR, leads to the formation of
the deamidated metabolite, NaAD (Trammell et al. 2016a). This could potentially be explained
by a bottleneck at the enzyme NADSYN, caused by exogenous treatment with NMN or NR.
Alternatively, bacteria possess deamidase enzymes such as the NMN deamidase PncC
(Gazzaniga et al. 2009), absent in mammals, which may contribute to the formation of NaAD
observed by Trammell and colleagues, by deamidating NMN and forming NaMN, the
precursor to NaAD in the Preiss-Handler pathway (Figure 5.4).
Our hypothesis proposed that a bacterial enzyme-mediated NMN deamidation step may
be involved in the formation of the deamidated metabolite, NaAD, following oral
administration with the amidated NAD+ precursor, NR. To investigate this hypothesis, we
specifically designed the labelled isotopes of NMN so that 15N labelling at both the pyridine ring and amide group of the nicotinamide moiety of NMN was achieved. As described in
Section 5.3.1 and illustrated in Figure 5.4, NMN deamidation forming NaMN, followed by the reamidation of NaAD into NAD+, would result in the loss of a single mass unit from the 15N
amide being removed. This would result in the conversion of M+7 NMN into M+6 NaMN via
PncC, formation of M+6 NaAD via NMNAT 1-3 (Berger et al. 2005) and final synthesis of
M+6 NAD+ via NADSYN (Hara et al. 2003) (Figure 5.4). Further, the appearance of M+6
NaAD would provide evidence for NMN or NR deamidation, rather than a bottleneck in
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Chapter 5 Host-microbiome interactions in NMN metabolism endogenous NAD+ synthesis at the enzyme NADSYN. This strategy would allow us to test our theory that the microbiome contributes to NMN deamidation, potentially via enzymes such as
PncC. Antibiotic treatment ablates bacterial species, and hence their deamidation enzymes, and subsequently we would expect a reduction in the abundance of M+6 labelled species detected in deamidated metabolites.
In this study, a small degree of M+6 labelled NaMN (Figure 5.16 B), NaAD (Figure
5.16 C) and NAD+ (Figure 5.16 D) was detected in the metabolite extracts from intestinal
homogenates following administration with NMN2. The formation of M+6 labelled NaMN
and NaAD with exogenous NMN treatment may suggest that the direct deamidation of NMN
through bacterial enzymes in the gut, and biosynthesis of NAD+ through the Preiss-Handler pathway metabolites, is a possibility. In contrast to our hypothesis, however, an increase in
M+6 labelled NaAD and NAD+ was observed in antibiotic treated mice following
administration with NMN, as denoted by the red bars in Figure 5.16 C and D, rather than an
expected decrease. These data contradict our hypothesis that NMN is deamidated through
interaction with bacterial deamidase enzymes in the gut. Instead, the presence of bacterial
species seems to be limiting the availability of NMN (Figure 5.16 A and E) and generation of
NaAD (Figure 5.16 C and G) and NAD+ (Figure 5.16 D and H) in the intestinal tissue.
Alternatively, these results indicate the generation of deamidated metabolites may not be
occurring through bacterial enzymes in the gut at all, as barely any M+6 labelled NaMN and
NaAD was detected in intestinal homogenates from mice treated with NMN1 (Figure 5.16 F
and G). Rather, these results put forward the possibility of a mammalian deamidase capable of
direct NMN or NR deamidation. While this poses an important question as to the identity of
this enzyme, whether it will meaningfully contribute to overall NAD+ metabolism in the host
is uncertain, as only a negligible proportion of deamidated metabolites, NaMN and NaAD
(<1pmol/mg tissue), were detected in this study (Figure 5.16 B and C).
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Figure 5.16 Deamidation pathway in the intestinal tissue of antibiotic treated mice following administration with NMN2 or NMN1 and 15N-amide-glutamine (15N-gln). Five-week-old female C57BL/6 mice were supplemented antibiotics in their drinking water for 11 days before NMN2 administration (50 mg/kg) by gavage. The intestinal tissues were collected at the 4-hr time point, and labelled isotopologues were detected in metabolite extracts from tissue homogenates using LC-MS/MS in MRM mode. The hypothesis that bacterial enzymes in the gut play a role in NMN metabolism by generating deamidated metabolites such as NaMN and NaAD, was contradicted by observing an increase in these metabolites following antibiotic treatment as observed in (A-D) for NMN2 and (E-H) for NMN1, and NaMN was largely absent after (B) NMN2 or (F) NMN1 administration. This may suggest a mammalian NMN deamidase could exist, though the physiological relevance may be negligible due to the low picomolar amounts detected. Data were analysed by two- way ANOVA with Sidak’s multiple comparisons test and represent mean ± s.d. (n=3-4 mice per group). *p<0.05, **p<0.01, ***p<0.001.
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Chapter 5 Host-microbiome interactions in NMN metabolism 5.4 Discussion
While intravenous delivery of NMN results in intact assimilation into peripheral tissues such
as the liver, kidney, brain and muscle, recent studies have revealed that the oral delivery of
NMN results in its cleavage at the glycosidic bond into free nicotinamide (Liu et al. 2018).
This was attributed to cleavage activity in the liver, however, orally administered NMN must also survive the gastrointestinal environment where bacteria have diverse enzymes to utilise
NAD+ precursors for their own metabolism (Gazzaniga et al. 2009), prior to reaching host
tissues.
In this chapter, we showed that the availability of orally delivered NMN in the intestinal tissue is tightly limited by competition with the gut microbiome. While this does satisfy the hypothesis that the microbiome contributes to the metabolism of orally administered NMN in vivo, the increase in NAD+ metabolites observed with antibiotic treatment was not anticipated
(Figure 5.8-Figure 5.13). This conclusion was drawn based on the specific conditions of this
study which involved a relatively modest dose of NMN (50 mg/kg) and its effect on levels of
NAD+ metabolites detected in the intestinal and liver tissue at an acute time point (4 hours). In
this study, the complete labelled versions of NMN were not detected in mice plasma from the
NMN2 or NMN1 cohort, which was consistent with the findings from a study conducted by
Liu and colleagues who also did not detect labelled NMN (or NR) in circulation at any time
point (0, 5, 15, 45 and 135 minutes) after orally administering the same bolus dose of NMN
(50 mg/kg) (Liu et al. 2018). In a study conducted by Mills and colleagues NMN was detected
in the plasma, with an increase detected as early as 2.5 minutes and returning back to baseline
by 15 minutes, however, the orally administered dose of NMN was much higher, at 300 mg/kg
body weight (Mills et al. 2016). As such, it is conceivable that the gut microbiome could
contribute to a threshold effect for dosing, thereby exogenously administered NMN or NR may
only reach peripheral tissues at a dose beyond the amount needed to saturate the microbiome
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Chapter 5 Host-microbiome interactions in NMN metabolism (i.e. >50 mg/kg), and mass balance studies that incorporate the terminal excretion of NMN and
other NAD+ metabolites from the body will be crucial to establish this. Importantly, while these
results demonstrate a tight competition for the uptake of NMN into the intestinal epithelium,
they were in close alignment with the finding by Liu and colleagues that a substantial
proportion of orally delivered NMN is decomposed to free nicotinamide in the liver (Liu et al.
2018). This study adds to these findings, where in addition to detecting an abundance of M+2
(Figure 5.9 G) and M+1 labelled NAM (Figure 5.12 G) in the liver, M+2 and M+1 NAM
were also abundantly detected in the intestinal tissues of mice administered NMN2 and NMN1,
respectively (Figure 5.8 G and Figure 5.11 G). This suggests that the cleavage of NMN at its
glycosidic bond may occur before it reaches the liver, in the gastrointestinal tract. As the half-
life for NAD+ turnover in the small intestine (< 30 minutes) is faster than that in the liver (2
hours) (Liu et al. 2018), detecting these labelled NAM isotopologues in the intestinal tissue at
earlier time points and comparing them to their detection in the liver is crucial to confirm this.
The increase in M+2 NAD+ in the intestinal tissues (Figure 5.8 J) and livers (Figure
5.9 J) of NMN2 administered mice, at levels higher than M+7 NAD+ (Figure 5.8 F, Figure
5.9 F), suggests new NAD+ is synthesised from the salvage of M+2 NAM rather than directly
from M+7 NMN. These data are consistent with a previous report (Liu et al) suggesting that
orally delivered NMN and NR undergo rapid metabolism by the liver to release free NAM,
raising the question of whether it is necessary to use NMN, rather than NAM, to raise NAD
levels. This also raises the question as to whether NMN acts as a prodrug of NAM and whether
administering NAM directly may be more effective in increasing cellular NAD+ levels, though
these comparative studies have not been conducted but are urgently warranted. There are
however, limitations to the therapeutic use of NAM as it inhibits the sirtuin enzymes (Avalos,
Bever, and Wolberger 2005; Bitterman et al. 2002), whereas NMN administration at a
relatively high dose (300 mg/kg) in mice over a 12-month treatment period yielded no such
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Chapter 5 Host-microbiome interactions in NMN metabolism side effects (Mills et al. 2016). A recent study revealed that NMN has longer retention in the
body compared to NAM (Kawamura, Mori, and Shibata 2016), with lower urinary excretion
of NAD+ catabolites, suggesting NMN has longer NAD+-raising potential over NAM. Future studies comparing the metabolic changes and therapeutic effects of NMN and NAM are needed to confirm this.
This study was initially undertaken to address questions of how the delivery of the amidated NAD+ precursor, NR, the precursor to NMN, could lead to a spike in the formation
of a deamidated metabolite, NaAD (Trammell et al. 2016a). We hypothesised that a
deamidation step could be mediated by bacterial enzymes in the gut microbiome, contributing
to an integrated model of host-microbiome NAD+ metabolism that has been described for the deamidation of NAM into NA by the nicotinamide deamidase, PncA (Frothingham et al. 1996),
in which case a reduction in the production of M+6 labelled NaAD following antibiotic
treatment was expected following NMN2 administration in mice. Contrary to this hypothesis,
the reverse occurred where an increase in M+6 NaAD labelling was observed (Figure 5.16 C
& G). Even if the involvement of bacterial deamidases to generate deamidated metabolites were true, based on the very low detection of M+6 labelled NaAD, whether it will meaningfully contribute to overall biosynthesis and metabolism of NAD+ inside the cell is uncertain.
Similarly, in the NMN1 cohort, the deamidation of M+6 labelled NMN1 isotope followed by
the re-amidation of M+6 labelled NaAD to M+7 labelled NAD in the presence of M+1 labelled
15N-glutamine was hypothesised (Figure 5.2), however, the opposite effect of targeting
bacterial enzymes using antibiotic treatment was observed (Figure 5.16 H). These data,
therefore, do not support the hypothesis that the appearance of deamidated metabolites
following oral delivery of amidated NAD+ precursors is mediated by bacterial enzymes in the
gut, and the question of why exogenous NR treatment leads to the acute formation of NaAD
observed by Trammell and colleagues, remains unanswered. Nevertheless, the appearance of
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Chapter 5 Host-microbiome interactions in NMN metabolism NaAD levels following supplementation with NR is consistently observed (Remie et al. 2020)
and is emerging as a sensitive biomarker for increasing intracellular NAD+ in cells and tissues.
While one reason for the appearance of NaAD could be from the deamidation of excess NAD+
as a way for cells to maintain a narrow range of cellular NAD+ concentration to regulate NAD+
homeostasis (Hara et al. 2019), this is unlikely to be the case as the amidation of NaAD into
NAD by NADS is an ATP dependent process, and the reversibility of this reaction is
thermodynamically unfavourable. Another possibility is that NaAD may be formed from
dephosphorylation of NAADP, a secondary messenger that plays a role in intracellular calcium
mobilisation, though the enzymes responsible for this proposed step are also yet to be
identified. Whether the formation of deamidated metabolites, NaMN and NaAD, can occur
through non-enzymatic acid-catalysed hydrolysis in the stomach or by another non-specific
deamidase enzyme is unknown. The redox state of NAD dictates its pH sensitive hydrolyisis,
with the reduced form NADH susceptible to degradation under acidic conditions but resistant
to high pH, and the inverse for NAD+ (Lowry, Passonneau, and Rock 1961). The products of
this acid-mediated degradation are not known to include deamidated products, and it is unclear
whether the pH of the stomach could influence the deamidation of NMN. Comprehensive
bioinformatic analysis of the entire NMN deamidase family, however, has revealed that NMN
deamidases are mainly found in bacteria, with a few examples in fungi (Hughes and
Williamson 1953; Sánchez-Carrón et al. 2013) and studies concerning deamidase activity in
mammals has been associated with microorganisms found in the stomach (Shimoyama et al.
1971).
Given our clear evidence for the preferential utilisation of orally delivered NMN by the
microbiome prior to host uptake, future studies should aim to characterise changes to the
microbiome itself during dosing with NAD+ precursors. Given our growing understanding of
the role of the gut microbiome in health, it is conceivable that physiological benefits observed
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Chapter 5 Host-microbiome interactions in NMN metabolism in preclinical models of ageing (Rajman, Chwalek, and Sinclair 2018) could, in part, be
mediated by changes to the microbiome. Overall, our findings for the role of the gut
microbiome to NMN metabolism will be important to the ongoing clinical development of
NAD+ precursors as drug candidates, with important implications for dosing and strategies
concerning their route of administration.
Study limitations
NAD+ metabolism in microbial organisms has been well-characterised and used as a guide to elucidate mammalian NAD+ metabolism (Gazzaniga et al. 2009). The idea that the gut
microbiome plays a role in the metabolism of NAD+ precursors is not unexpected, however most animals studies that use NAD+ precursors, such as NMN and NR, fail to take this into
consideration. The preliminary findings from this study provide valuable insight into the role
of the gut microbiome in the metabolism of NAD+ precursors such as NMN. In future, it will
be important to consider whether the route of administration and dose of NAD+ precursors in
preclinical and clinical studies is appropriate.
A higher dose may not necessarily translate to improved efficacy or superior therapeutic
outcomes, in fact, a single dose of NMN at 62.5 mg/kg had the strongest protective effect
against ischemic brain injury in mice compared to the higher doses administered at 125, 250
and 500 mg/kg in mice (Park et al. 2016). In another study, long term administration of NMN
at a lower dose (100 mg/kg/day) was better at improving functional parameters such as oxygen
consumption, energy expenditure and physical activity, whereas the higher dose (300
mg/kg/day) was associated with greater improvements in tear production, bone density and
myeloid-lymphoid composition (Mills et al. 2016), indicating that the optimal dose of NMN
may differ significantly depending on the physiological target. A small bolus dose of NMN at
50 mg/kg was administered to mice in this study which may have been directly consumed by
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Chapter 5 Host-microbiome interactions in NMN metabolism microbial organisms in the gut, accounting for the low levels of NAD+ metabolites detected in
peripheral tissues. Future studies investigating NMN metabolism should aim to test a wider
range of doses to establish whether a higher dosing can overcome the potential threshold effects
of the microbiome on host availability of orally administered NAD+ precursors.
One major limitation of this study is that the small (duodenum, proximal jejunum,
middle jejunum, distal jejunum, and ileum) and large (cecum, ascending colon, and descending
colon) intestinal segments were combined, causing a loss of functional heterogeneity. Different
segments not only harbour different populations of microbes, but also identify with different
cell types such as enterocytes, dendritic cells, neuroendocrine cells, goblet cells, proliferating
stem cells, Paneth cells, immune cells, muscle cells, neurons and adipocytes (Kong, Zhang,
and Zhang 2018; Zhang et al. 2018a; Zhao et al. 2015). The unique functions and differences
in metabolism of these cell types along the length of the intestines could significantly vary and
performing metabolite analyses in separate intestinal segments will provide further insight into
this.
Another limitation is that the changes to the NAD+ metabolome in this study were
examined at a single time point, rather than over a time course. The 4-hour time point chosen
in this study may have been too long to examine the minute-order kinetics of NMN transport into the cell, as previously reported (Mills et al. 2016; Yoshino et al. 2011). Future studies should aim to sample at tighter and more frequent time points to account for the minute- and hourly-order kinetics of NMN transport and metabolism.
The high variability in the intestinal concentrations of metabolites between NMN2
(Figure 5.8) and NMN1 (Figure 5.11) cohorts may be attributed to the sample measurements being acquired in separate LC-MS batches. In addition to the natural biological variance between experimental mice, slight differences in HPLC column conditions over time and/or
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Chapter 5 Host-microbiome interactions in NMN metabolism sample processing periods may change metabolite signals, contributing to the variability seen
between intestinal samples across separate cohorts. Further, the standard calibration curves
used to calculate the molar amounts of NAD metabolites were generated for each sample batch
and included internal standards however, were prepared in LC-MS-grade water rather than the sample matrix which consists of components that may cause ion suppression, a significant reduction in analyte signal due to competing or interfering matrix components with the metabolite of interest (Annesley 2003; Stokvis, Rosing, and Beijnen 2005). While standard calibration curves and samples were all normalised to the internal standard (thymine-d4), the difference between the internal standard values in standards and samples was on average approximately 7-fold higher in the NMN2 cohort and 10-fold higher in the NMN1 cohort, suggesting a potential matrix effect on ionisation efficiency. This limitation should be addressed in future studies by performing experiments which test the degree of ion suppression
(Annesley 2003), determine the conditions that influence ionisation efficiency (such as type of sample matrix, extraction method and buffers, or structure of the metabolite of interest) and develop ways to minimise these interferences to achieve greater accuracy between batches.
One of the main aims of this study was to explore the mechanism of NMN transport into the cell using labelled isotopes of NMN. This assumed the presence and function of transporters SLC12A8 and ENTs (SLC29A1 and SLC29A2) as well as enzymes CD73 and
NRK1/2, however this was not experimentally demonstrated. This should be addressed in future studies through genetic deletion of SLC12A8 (Grozio et al. 2019a), NRK1/2 (Ratajczak et al. 2016) or supplementation with small molecule inhibitors such as adenosine-5′ -(α ,β - methylene)-diphosphate (AOPCP) which inhibits CD73 activity (Bhattarai et al. 2015) and dipyridamole (DIPY) or S-(4-Nitrobenzyl)-6-thioinosine (NBTI) which inhibit plasma membrane nucleoside transporters such as ENTs (Jarvis 1986; Soriano-Garcia et al. 1984).
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Chapter 5 Host-microbiome interactions in NMN metabolism Finally, to consolidate the hypothesis that the gut microbiome acts in competition with host intestinal tissues for the uptake of orally delivered NMN, the measurement of NAD+ metabolites in the faecal contents of mice will provide better insight into whether it is being properly absorbed in the gut and metabolised, rather than being excreted.
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Chapter 6 General Discussion
General Discussion
It has been over 110 years since NAD+ was first discovered as an essential redox cofactor
required for yeast fermentation, and its biological significance has since grown to include other
roles including as a critical substrate for proteins regulating DNA repair, metabolism and
longevity (Katsyuba et al. 2020). Declining NAD+ levels with advanced age are implicated in
age-related diseases (Camacho-Pereira et al. 2016; Gomes et al. 2013; Massudi et al. 2012a),
compromising cellular redox reactions and the activity of enzymes such as the sirtuins, PARPs
and CD38/CD157, which consume NAD+ as a co-substrate (Braidy et al. 2011b; Camacho-
Pereira et al. 2016; Frederick et al. 2016; Pang et al. 2015; Schultz and Sinclair 2016; Wang et
al. 2013; Wilk et al. 2020; Yaku, Okabe, and Nakagawa 2018a). In a bid to halt the decline in
NAD+ levels, supplementation with NAD+ precursors such as NMN and NR has show potential to improve healthspan and lifespan (Bertoldo et al. 2020; Das et al. 2018; de Picciotto et al.
2016; Diguet et al. 2018; Mills et al. 2016; Tarantini et al. 2019; Trammell et al. 2016c;
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Chapter 6 General Discussion
Yoshino et al. 2011; Zhang et al. 2016a) with clinical studies underway (Airhart et al. 2017a;
Dolopikou et al. 2019; Elhassan et al. 2019; Irie et al. 2020; Martens et al. 2018; Remie et al.
2020).
Despite the popularity of this strategy, key details of how these supplements raise
NAD+ levels in the body are missing, which may lead to more effective ways to increase their
therapeutic potential. Immediately prior to submission of this thesis, a new pathway for NAD+
biosynthesis was revealed. This involved the reduced form of NR (NRH) (Giroud-Gerbetant et
al. 2019; Yang et al. 2019) acting as a substrate for the enzyme adenosine kinase (AK) (Yang
et al. 2020). Critically, the action of this pathway allows for NRH treatment to increase NAD+
levels up to 10-fold in mammalian cell lines and mouse models (Giroud-Gerbetant et al. 2019;
Yang et al. 2019; Yang et al. 2020), compared to the 2-4 fold increases in NAD+ levels achieved by other precursors such as NMN and NR (Trammell et al. 2016b; Yoshino et al. 2011). The limited increase in NAD+ levels with NMN or NR treatment aligns with other arguments that
NAD homeostasis is tightly maintained, even when the supposed rate-limiting enzyme
NAMPT is over-expressed (Hara et al. 2019). Understanding how this new arm of NAD+
synthesis involving NRH and AK escapes this regulatory constraint, and importantly NAD
metabolism in general, will have important therapeutic implications.
Our work was motivated by the fact that in clinical trials, the effects of treatment with
NAD+ precursors have been mixed. While NR is well tolerated at high doses of up to 2 grams
per day, only mild benefits on blood pressure and exercise performance were demonstrated in
elderly individuals (Dolopikou et al. 2019; Martens et al. 2018). We speculate that this underwhelming results is due to the fact NAD+ precursors are commonly administered via the
oral route and must persist in the gastrointestinal tract to deliver their therapeutic payload
(Airhart et al. 2017a; Elhassan et al. 2019; Irie et al. 2020; Remie et al. 2020; Trammell et al.
2016b). Differences in the gut microbiome of test subjects may be a source of variation in
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Chapter 6 General Discussion
results. . Currently, the contribution or effect of the microbiome has on the metabolism and
bioavailability of these compounds to the host is unknown. For example, whether the
microbiome plays a role in the appearance of deamidated metabolites such as NaAD and
NaMN following oral administration with the amidated precursor NR (Elhassan et al. 2019;
Trammell et al. 2016a) has neither been confirmed nor ruled out. The studies conducted in this
thesis sought to achieve a better understanding of the gut microbiome interactions with host
metabolism of the NAD+ precursor NMN. Observations of E. coli rapidly catabolising NMN
in the periplasmic space (Chapter 4) and antibiotic treatment increasing NMN uptake via NR
at the gut epithelium (Chapter 6) convinced us that the gut microbiome acts in competition with
the host for orally supplemented NMN. The conclusions drawn from these preclinical findings
demonstrate the importance of reconsidering the route of administration and dosing
requirements in the therapeutic use of NAD+ precursors.
6.1 Quantifying the NAD metabolome using LC-MS/MS—So close, yet [not] so far
At the start of this thesis program, LC-MS-based methods using multiple-reaction-monitoring
(MRM) to quantify the NAD+ metabolome were limited. There was no method on hand that
could simultaneously measure the different NAD metabolites and their isotopologues. The
latter is needed to trace how NMN is assimilated and to reveal the pathways by which NMN is
transported into the cell and utilised in vivo. A crucial first step in this thesis was to establish a
reliable LC-MS/MS method to quantify how isotope labelled material is propagated through
the NAD metabolome. A key challenge was achieving LC separation of the acidic metabolites
NaAD and NaMN from their amidated counterparts NAD+ and NMN, as the difference in mass
shift of only a single Dalton in mass can confound downstream analyses. Ultimately, we
developed and validated an LC-MS approach that can be used to quantify metabolism of NMN
that is isotopically labelled. This method can be applied to biological samples from in vitro and
in vivo models.
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Chapter 6 General Discussion
The quantification of NAD metabolites has come a long way, but progress has been
most rapid in recent years. Nearly two decades ago, Evans and colleagues were the first to
report an LC-MS/MS method for the measurement of NAD+ in cellular extracts (Evans et al.
2002). They used four stable isotope labelled precursors tryptophan (Trp), quinolinic acid
(QA), nicotinic acid (NA) and nicotinamide (NAM) to measure the incorporation of labelled
moieties into NAD+ but no other NAD+ metabolites were quantified. Further, an ion trap mass
spectrometer was used which requires each selected reaction monitoring (SRM) transition to
be scanned individually, rather than a triple-quadrupole mass spectrometer which uses MRM
to measure multiple metabolites of interest in a single injection. Shortly after, Yamada and
colleagues improved NAD+ quantification using a triple-quadrupole mass spectrometry
method and were the first to directly measure NAD+ metabolites NMN and NaMN in any cell
type. They were however, limited by a subset of NAD+ metabolites where a majority of the metabolites (NADH, NADP+, NADPH, QA, NR, NaR, NA, NAM and NaAD) were below the
detection limit (Yamada et al. 2006). In 2010, Evans and colleagues developed an LC-MS/MS
based assay with hydrophilic interaction liquid chromatography (HILIC) using a Luna NH2
column which provided good retention and chromatographic resolution of a defined core set of
NAD+ metabolites (Evans et al. 2010). Following this, Trammell and colleagues developed a
highly reliable and robust LC-MS/MS method to quantify a total of 19 NAD+ metabolites
including all major metabolites involved in NAD+ biosynthesis pathways (NAM, NMN,
NAD+, NR, NA, NaR, NaMN and NaAD) (Trammell et al. 2016a). The only limitation was the
need for two separate extraction methods, acidic and basic, to accurately quantify NAD+
metabolites.
The LC-MS/MS method developed in this thesis (Chapter 3) was not adapted from the
aforementioned methods specialising in the detection of NAD metabolites, but instead from a
metabolomics protocol by Yuan and colleagues that targeted more than 250 compounds and
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covered all major metabolic pathways including glycolysis, the TCA cycle and the pentose
phosphate pathway (PPP) (Yuan et al. 2012). By using a type of HILIC-based chromatography,
i.e., amide phase, to retain extremely polar compounds, we achieved good resolution of the
eight core NAD+ metabolites (NAM, NMN, NAD+, NR, NA, NaR, NaMN and NaAD) in a
sensitive and reproducible manner. The strongest advantage of the method developed here was
the use of a triple quadrupole mass spectrometer in MRM mode to detect a total of 64 individual
isotopologues each with unique MRM (Q1>Q3) transitions, all in a single injection for each
sample. While there were challenges with the initial separation of the acidic metabolites NaAD
and NaMN to their amidated counterparts NAD+ and NMN respectively, we overcame this by
adjusting the pH of our mobile phase buffer from pH 9.4 to pH 5 using acetic acid and
developing deconvolution scripts to resolve the overlapping pairs of closely related metabolites using MATLAB software.
This method was then validated in primary hepatocytes isolated from mice and in
C2C12 and HEK293 mammalian cell lines to show the successful quantification of both unlabelled and labelled NAD+ metabolites can be detected in a biologically relevant in vitro model. This also served as a validation of our isotope labelling strategy, confirming that NMN is metabolised in mammalian cells via the canonical pathway, as expected. This involved the direct synthesis of NAD+ via NMNAT and recycling of NAM, the by-product of NAD+-
consuming activities through the salvage pathway, as observed by increases in labelled NAD+
and NAM isotopologues in cell lysates following supplementation with labelled NMN
isotopes.
In recent years, LC-MS-based metabolomics have become one of the most reliable and
sensitive techniques to quantify the NAD+ metabolome and further, to study NAD+ precursor
metabolism and transport into mammalian cells (Bustamante et al. 2018; Grozio et al. 2019a;
Liu et al. 2018; Ratajczak et al. 2016; Trammell et al. 2016a). Despite uncertainties around
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Chapter 6 General Discussion
which biological mechanisms are most important for mediating the benefits of elevating NAD+
levels, the importance of rigorous measurements of NAD+ and its metabolites are increasingly
recognised. For example, the consistent appearance of elevated NaAD levels in peripheral
tissues following the administration of the NAD+ precursor NR under preclinical and clinical
conditions (Remie et al. 2020; Trammell et al. 2016a) suggest there are aspects of NAD+
metabolism which are not understood, which may have gone unnoticed without their
serendipitous observation using LC-MS-based techniques. Future studies will require further
evaluation of these pathways and additional isotope tracer studies that employ a diverse range
of biological systems are needed.
6.2 Mechanisms of NMN uptake in gut epithelial tissue — ¿Por qué no los dos?
Whether the gut epithelium takes up NMN intact has been a highly debated question in this
field. The dilemma was whether the direct or indirect route is the most efficacious absorption
route for NMN, with important implications for which NAD+ precursor has greater efficacy.
Ultimately, our results added weight to the idea that NMN is taken up indirectly as NR. Our
findings highlighted the role and physiological importance of NR in raising NAD+ levels in the cell.
Briefly recapping that ongoing debate, recent work argued that the fast pharmacokinetics of exogenously supplemented NMN is due to direct and intact transport of
NMN, where it is immediately utilised for NAD+ biosynthesis (Mills et al. 2016; Yoshino et
al. 2011). This led to the hypothesis that there is an effective NMN transporter, which remained
elusive for several years until a recent paper characterised Solute Carrier Family 12 Member 8
(SLC12A8) as a dedicated NMN transporter (Grozio et al. 2019a). The interpretation of this
work has been challenged, due to differences in the bioanalytical method used to detect these
metabolites (Schmidt and Brenner 2019), with counter-arguments (Grozio et al. 2019b) that
have fuelled this debate. A competing idea argues that NMN transport is indirect and requires
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Chapter 6 General Discussion
dephosphorylation to NR prior to uptake, where it is re-phosphorylated by NR kinases (NRK1
or NRK2) to form NMN, as a rate-limiting step required for the utilisation of exogenous NMN
to raise intracellular NAD+ levels (Ratajczak et al. 2016).
Using isotope labelled NMN, the results presented in this thesis supported the indirect
mechanism for NMN transport into the cell involving the dephosphorylation of NMN to NR.
This was based on the significant detection of intact labelled NR following supplementation
with labelled NMN in extracts from primary hepatocytes, C2C12 and HEK293 cells (Chapter
3), supernatants of E. coli bacteria (Chapter 4) and intestinal and liver tissues of mice (Chapter
5). These findings are consistent with previous tracer studies in the human hepatocellular
carcinoma (HepG2) cell line where treatment with 18O-NMN labelled at the carboxamide
oxygen of the nicotinamide moiety resulted in the appearance of 18O-NR in the conditioned
media, parallel to the disappearance of 18O-NMN, concurrent with steady intracellular
enrichment of 18O-NR. These results support the mechanism by which exogenous NMN first
undergoes dephosphorylation to NR prior to entry into the cell (Ratajczak et al. 2016).
Our results did not entirely exclude the possibility that NMN may still be transported
intact and directly via a dedicated NMN transporter, as intact labelled NMN was detected at
earlier time points in both C2C12 and HEK293 cell lysates (Chapter 3). In future work, it may
be interesting to determine whether there are conditions under which utilisation of one
mechanism over another may be favoured in the context of treating pathological conditions or
age-related diseases. For example, a recent study reported that high expression levels of the
NMN transporter SLC12A8 and its genetic polymorphism leads to a better prognosis for
patients with pancreatic ductal adenocarcinoma (Feigin et al. 2017) and breast cancer (Kim et al. 2018) respectively. Furthermore, SLC12A8 expression is upregulated in aged ileum of mice and supplementation with NMN could maintain NAD+ in this tissue at levels comparable to
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Chapter 6 General Discussion
those of young mice (Grozio et al. 2019a). These results highlight that NMN and its transporter
SLC12A8 could be used as potential new therapeutic targets in the treatment of certain cancers.
One argument to support the direct NMN transporter mechanism shown in previous studies is in NRK1-deficient mice, intraperitoneal injection of NMN still increased NAD+
levels in the liver, kidney, brown adipose tissue and skeletal muscle, though the statistical
significance of these results was not shown. An alternative explanation for these results could
be that expression of NRK2 is highly restricted to skeletal muscle and could compensate for
the loss of NRK1 to re-phosphorylate NR to NMN in this tissue (Ratajczak et al. 2016).
Another study treated primary myotubes from NRK1/NRK2 double knockout mice with
unlabelled NMN, with increased NAD+ levels compared to controls, though again, significance
was not reported (Fletcher et al. 2017). The strongest argument supporting the existence of
direct NMN transport is that the minute-order uptake of NMN reported in previous studies
(Grozio et al. 2019a; Mills et al. 2016; Yoshino et al. 2011) cannot be explained by the hourly
kinetics of NMN to NR conversion, which can occur over the course of 24 hours (Ratajczak et
al. 2016). Nonetheless, these nascent findings suggest the ability to raise NAD+ levels from
exogenous NMN may not be entirely dependent on its conversion to NR prior to entry, and its
assimilation into the cell via the NMN transporter may be of equal physiological importance.
6.3 Host-microbe interactions in NAD+ precursor metabolism—Friend or foe?
The results of this investigation were surprising. While we established that the gut microbiome
did play a major role in the metabolism of orally administered NMN, it was not in the direction of boosting NAD+ metabolism, as initially hypothesised. Our discovery will be an important contribution to determining which route may be most effective for delivering NAD+ precursors.
It was suggested that high level of expression of SLC12A8 in the duodenum, ileum and jejunum allows the gut to directly transport NMN (Grozio et al. 2019a). In this study, intact
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Chapter 6 General Discussion
NMN was detected in the intestinal and liver extracts of antibiotic-treated mice only (Chapter
5). This providing evidence that the gut microbiome may limit NMN bioavailability following a bolus dose. Reducing the bacterial load in the gut translated to a greater abundance of most
NAD+ metabolites in host tissues, indicating that this impact is systemic. Remarkably, antibiotic-treated mice that did not receive a bolus of NMN still showed more abundant NAD+ metabolites. From this, we speculated that the competition between the host and microbiome is not only for NMN, but also for NA and NAM, which may be obtained from background dietary sources.
The above interpretation does not take into account the effect of antibiotic treatment on the integrity of the intestinal epithelium, as the gut microbiome is important to support the renewal of the epithelial lining, maintain optimal barrier permeability and protect against epithelial cell injury (Kang et al. 2013a; Mathewson et al. 2016; Stefka et al. 2014; Zhan et al.
2013). Methods to test gut permeability in vivo, for example by orally administering non- digestible tracer molecules and quantifying them in the urine or blood of antibiotic-treated mice, will be crucial to establish this (González-González et al. 2019).
We acknowledge that the lack of NMN detected in our studies may reflect the relatively late timing of sampling, where sampling at four hours would have missed minute-order transport of NMN into the cell as indicated in previous studies (Yoshino et al. 2011; Mills et al. 2016). With this in mind, caution is needed in the interpretation of these data. Time course experiments were not an option for us due to limited availability of our isotope labelled NMN
. The timing of sampling (4 hours) was based on previous studies that detected a peak in the deamidated metabolite NaAD in the peripheral tissues of NR-administered mice (Trammell et al. 2016b). Our data and those from others show that the NAD+ precursors NAM, NA and NR have unique pharmacokinetic profiles, where the same metabolite peaked at different times following oral administration (Trammell et al. 2016b), highlighting the importance of
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Chapter 6 General Discussion analysing samples over a time course, rather than a single time point. Future studies should address these study limitations by sampling at tighter and more frequent time points to address the potential pharmacokinetic influences on NMN transport mechanisms.
Likewise, spatial issues could explain our inability to detect NMN. We analysed a bulk sample of the whole length of the small intestine, rather than separating and analysing different intestinal segments (duodenum, ileum, jejunum, cecum and colon). These subsections differ in the density and diversity of bacterial species, with likely differences in the metabolism of
NAD+ precursors via different routes (Gazzaniga et al. 2009). Further, expression of the putative NMN transporter SLC12A8 is not uniform across the entire intestine, and a homogenous tissue lysate may have missed these differences.
6.4 Preliminary evidence for host NMN deamidation—it’s not you, it’s me
Our investigations here were based on the hypothesis that the appearance of deamidated NAD+ intermediates NaMN and NaAD following oral administration with NR (Trammell et al.
2016b) is mediated by bacterial NMN deamidase enzymes in the gut, such as PncC. Consistent to our hypothesis, we showed that mammalian cells in culture lacked the ability to deamidate
NMN because the levels of labelled NaMN and NaAD detected in our in vitro studies were negligible (Chapter 3). In parallel, we detected labelled NaMN in E. coli culture media (Chapter
4), which indicated the conversion of NMN to NaMN by intracellular PncC and NaMN export by an unknown mechanism (Galeazzi et al. 2011). Based on these in vitro results, we concluded the conversion of NMN to NaMN is dependent on the action of bacteria. During preparation of this thesis, an elegant study by Shats and colleagues showed that orally delivered
NAM undergoes deamidation into NA by bacterial deamidase enzymes in the gut (Shats et al.
2020). This study used a combination of deuterated NAM, antibiotic treatment and the repopulation of germ-free wild-type mice with wild-type and pncA knockout E. coli to
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Chapter 6 General Discussion
ultimately confirmed PncA-mediated conversion of NAM to NA is drives the appearance of
deamidated NAD+ metabolites in host tissues.
Interestingly, our in vivo experiments yielded different results. We found that labelled
NMN deamidated into NaMN in antibiotic treated mice, but not the control group (Chapter 5,
Figure 5.14). Validation is still required because the amount of NaMN and NaAD detected
were low compared to the amidated metabolites NR, NAM, NAD+ and NMN. A crucial difference between this study and that of Shats el al (2020) is the use of NMN rather than NAM, with differences in the route of uptake that could explain this discrepancy. In these results, when the gut microbiome was ablated, host tissues were exposed to a greater concentration of
NMN, and it is quite plausible that NMN deamidation pathways were engaged under this circumstance. This means that although NAM deamidation is definitely bacterial dependent, whether mammalian cells are incapable of NMN deamidation remains to be addressed.
Importantly, this work may contribute to understanding the changes that occur in microbial and host NAD+ metabolism—separately and as a whole—following supplementation with NAD+
precursors.
6.5 Beyond NAD metabolism—blue-sky thinking
Beyond their potential role in the metabolism of NAD+ precursors, the microbiome is well-
known to contribute to the metabolism of xenobiotics including commonly used compounds
such as metformin, gemcitabine, oxaliplatin and levodopa, which are important for the
treatment of type II diabetes, cancer and Parkinsons’s disease, respectively (Clarke et al. 2019;
Kang et al. 2013b; Pryor et al. 2020). This can have important clinical impacts.
A recent work by Rekdal and colleagues demonstrated the detrimental impact of the microbiome on xenobiotic compounds. This group uncovered fundamental pathways for the microbial metabolism of levodopa (L-dopa), an orally administered drug in the treatment of
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Chapter 6 General Discussion
Parkinson’s disease (PD) (Rekdal et al. 2019). Microbial metabolism of L-dopa was associated
with reduced efficacy of the drug, high interpatient variability and development of adverse side
effects. Thus understanding these mechanisms and host-microbe interactions has the potential
to enhance the development of L-dopa in the treatment of PD, and it is conceivable that a
similar phenomenon may be at play in the case of therapeutic administration of NAD
precursors.
On the opposite end of the spectrum, we also see beneficial impacts for the the
microbiome in mediating the benefits of drug treatments. A recent paper suggested the
therapeutic effect of the anti-diabetic drug metformin may be due to changes in gut microbial
diversity and abundance (Pryor et al. 2020), with improvements The in microbiota-induced
maintenance of intestinal barrier integrity, and regulation of the secretion of gut hormones
involved in appetite control.
Given the biological significance of NAD+ in microbial systems, with more pathways
and enzymes involved in its biosynthesis than in mammals (Gazzaniga et al. 2009), it is
conceivable that the benefits of NAD+ precursors are in part mediated by complex host-microbe
interactions. Whether these interactions benefit or limit the efficacy of orally administered
NAD+ precursors remain to be seen, and may have important clinical implications for the use of NAD+ precursors in the treatment of age-related disease.
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Appendix
Figure A.1 Differential expression analysis of combined bacterial taxonomic levels in NMN2 cohort. Panels show MA-plots for differential testing between treatment groups. Y- axis shows the log2-fold change of the OTU. X-axis shows the mean of normalised counts of the OTU. Differential OTUs from treatments were tested using the Wald test in the R package, DESeq2, with red coloured points significant at an adjusted p.value < 0.05 using the Benjamini- Hochberg correction for multiple testing.
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Figure A.2 Differential expression analysis of combined bacterial taxonomic levels in NMN1 cohort. Panels show MA-plots for differential testing between treatment groups. Y- axis shows the log2-fold change of the OTU. X-axis shows the mean of normalised counts of the OTU. Differential OTUs from treatments were tested using the Wald test in the R package, DESeq2, with red coloured points significant at an adjusted p.value < 0.05 using the Benjamini- Hochberg correction for multiple testing.
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APPENDIX
Nicotinamide mononucleotide (NMN) deamidation and indirect regulation of the NAD metabolome
Lynn-Jee Kim1, Timothy J. Chalmers1, Greg C. Smith1, Abhirup Das1, Eric Wing Keung Poon2, Jun Wang2,3, 4 1,5 6* 1* Simon P. Tucker , David A. Sinclair , Lake-Ee Quek , Lindsay E. Wu
1School of Medical Sciences, UNSW Sydney, NSW 2052, Australia
2GeneHarbor (Hong Kong) Biotechnologies Limited, Hong Kong Science Park, Shatin, Hong Kong SAR, China
3School of Life Sciences, The Chinese University of Hong Kong, Hong Kong SAR, China
4Jumpstart Fertility Pty Ltd, Melbourne, Victoria, Australia
5Harvard Medical School, Boston, MA, United States of America
6School of Mathematics and Statistics, The University of Sydney, NSW 2007, Australia
*These authors contributed equally to this work. Correspondence: [email protected] +61 2 9385 1621
ABSTRACT
Treatment with nicotinamide mononucleotide (NMN) is a prominent strategy to address the
age-related decline in nicotinamide adenine dinucleotide (NAD+) levels for maintaining
aspects of late-life health. It is assumed that exogenous NMN is directly incorporated into the
NAD+ metabolome in mammals via the canonical recycling pathway. Here, we show that NMN
can undergo direct deamidation and incorporation via the de novo pathway, which is in part
mediated by the gut microbiome. Surprisingly, isotope labelling studies revealed that
exogenous NMN treatment potently increased the endogenous production of unlabelled NAD
metabolites, suggesting that exogenous NMN impacts the NAD metabolome through indirect
means, rather than through its direct incorporation. This included a striking increase in
endogenous production of the metabolites nicotinic acid riboside (NaR) and nicotinamide
riboside (NR) which was amplified in antibiotics treated animals, suggesting the production of
endogenous NaR/NR through altered metabolic flux, enzyme kinetics and/or an as-yet unidentified pathway that interacts with the gut microbiome.
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Nicotinamide adenine dinucleotide (NAD+) is an essential redox cofactor central to metabolic
processes such as glycolysis, the tricarboxylic (TCA) cycle and fatty acid oxidation1,2. NAD+
is also consumed by enzymes such as the sirtuins3 and poly(ADP-ribose) polymerase (PARP) enzymes4 which are mediators of genome stability5 and DNA repair6. Given the essential role
of this metabolite, the decline in NAD+ that occurs during biological ageing7-12 and disease
states13-15 has gained attention as a target for therapeutic intervention16. Strategies to boost
NAD+ levels through supplementation with NAD precursors such as nicotinamide
mononucleotide (NMN) and nicotinamide riboside (NR) are emerging as promising
therapeutics12,16-23. Historically, dietary supplementation with the NAD precursors nicotinic acid (Na) or nicotinamide (Nam) was used to prevent chronic NAD deficiency, which causes pellagra. When these micronutrients are replete, the step converting Nam into NMN by the enzyme nicotinamide phosphoribosyltransferase (NAMPT) is rate limiting in NAD synthesis24,
and the use of NAD precursors that occur after this step, namely NMN and NR, have gained
prominence as a strategy to raise NAD+.
One surprising aspect of this strategy is the striking appearance of the deamidated metabolite nicotinic acid adenine dinucleotide (NaAD) following oral delivery with the amidated metabolite NR25. NR is phosphorylated into NMN by NR kinases (NRK1/2)26,27, and then
adenylated into NAD+ by NMNAT enzymes (NMNAT1-3)28-33, effectively bypassing NaAD,
which is an intermediate of the Preiss-Handler or de novo pathway34,35. In contrast, bacteria
have a well-characterised NMN deamidase enzyme, PncC36 that prevents the accumulation of
NMN, which inhibits the bacterial DNA ligase37-39. One theory to explain the increase in NaAD
with NR treatment25 could be that NMN and NR assimilation follows a non-canonical route
that combines steps of both microbial and mammalian processes, whereby NMN is deamidated
into NaMN or NaR prior to its uptake into mammalian tissue, and then assimilated into NAD+
via the intermediate step of NaAD. This could explain the appearance of NaAD following NR
216 APPENDIX
supplementation25, however an unexplained aspect is that the delivery of labelled NR results
in the formation of unlabelled NaAD25.
Here, we use targeted metabolomics to trace the in vitro and in vivo metabolism of strategically designed NMN isotopologues to answer these questions. We show that NMN can be incorporated following its deamidation and metabolism via the de novo route, which is in part mediated by the microbiome. We further show that ablation of the microbiome by antibiotic treatment increases the uptake and conversion of orally delivered NMN into the NAD metabolome, and that isotope labelled NMN overwhelmingly presents in intestinal tissue in the form of NaR and NR. Contrary to the assumption that exogenous NMN treatment raises NAD+
levels solely through its direct incorporation into the NAD metabolome, we show that treatment
with isotope labelled NMN increases the levels of endogenous, unlabelled NAD metabolites.
Overall, our results provide unique insights into the assimilation of orally delivered, exogenous
NMN into gastrointestinal tissue, and raise questions around how exogenous precursors alter
the NAD metabolome.
RESULTS
NMN treatment alters the de novo arm of NAD+ synthesis
According to canonical models of mammalian NAD homeostasis, the metabolism of NMN, an
amidated intermediate in the recycling pathway, does not intersect with the de novo pathway,
which utilises deamidated intermediates. Unlike mammals, bacteria present in the gut
microbiome do encode deamidase enzymes such as PncC, which deamidates NMN into
nicotinic acid mononucleotide (NaMN) for metabolism via the de novo pathway36. To test whether the gut microbiome alters the in vivo metabolism of orally administered NMN, we
used mice that were exposed to a course of antibiotics to ablate the gut microbiome (Supp. Fig.
1). These animals received a bolus of NMN (500 mg/kg) by oral gavage, and four hours later,
217 APPENDIX animals were sacrificed and tissues rapidly preserved for targeted metabolomic analysis (Fig.
1). We focused our analyses on the gastrointestinal tract (GIT) and the liver, as these two tissues have high levels of NAD synthetase (NADS) activity40, and are the primary sites of uptake and metabolism for orally delivered compounds. In agreement with previous work25, NMN treatment increased the abundance of the de-amidated metabolites NaR and NaMN in both the gastrointestinal tract (GIT) (Fig. 1a, b) and liver (Fig. 1d, e), while NaAD was increased in the liver (Fig. 1f), matching previous findings for NR25. Interestingly, this was completely abolished in antibiotic treated animals, where NMN treatment instead led to a spike in the amidated metabolites NR (Fig. 1g, j) and NMN (Fig. 1h, k), and abolished the increase in liver
NaAD (Fig. 1f). To highlight the inverse relationship between amidated and de-amidated metabolites during antibiotics treatment, the abundance of each deamidated metabolite was expressed as a ratio of its amidated counterpart (Fig. 1m-r), highlighting a profound role for the microbiome in dictating the roles of the de-amidated and amidated arms of NAD metabolism.
Strategic isotope tracing of NMN metabolism
We next sought to carefully test whether exogenous NMN was indeed undergoing direct deamidation prior to its incorporation into the NAD metabolome using isotope tracing studies.
We designed two separate isotopologues of NMN that were strategically labelled at positions that would answer our hypothesis of NMN deamidation. The first of these, designated as
NMN1, was 13C labelled at all five carbon positions of the ribose moiety for an M+5 mass shift, and 15N labelled at the pyridine ring for an overall M+6 mass shift (Fig. 2a, Supp. Fig.
2). In the last step of the de novo pathway, the enzyme NAD synthetase (NADS) amidates the carboxylic acid of NaAD using an ammonia intermediate derived from the amide group of glutamine, yielding glutamate (Fig. 2). By delivering the NMN1 (M+6) isotope in the presence of 15N-amide labelled glutamine (M+1), the presence of M+7 labelled NAD+ with an additional
218 APPENDIX
mass shift from the nicotinyl amide would indicate that the original amide N atom had been
lost during deamidation and replaced by the 15N amide from 15N-Gln, indicating incorporation
of NMN into NAD+ via prior deamidation and the de novo pathway (Fig. 2a). To complement
this experiment, we designed a second isotope, designated as NMN2, where all five carbons of
the ribose moiety were 13C labelled, and both the pyridine ring and primary amide positions
were 15N labelled, for an overall M+7 mass shift (Fig. 2b, Supp. Fig. 2). When delivered in a
separate experiment, if NMN2 (M+7) underwent deamidation prior to its incorporation, the
15N amide would be lost and replaced by an unlabelled amide from the endogenous glutamine
pool, resulting in M+6 labelled NAD+. By comparing the ratios of M+7 and M+6 labelled
NAD+ in each experiment, we could quantify the proportion of NMN that had been
incorporated into NAD+ following deamidation and assimilation by the de novo pathway. This
would be supported by comparing the ratios of M+1 and M+2 labelled Nam, which is released
by NAD+ consuming enzymes, however this interpretation would be complicated by the
recently described role of the bacterial nicotinamide (Nam) deamidase PncA in systemic
mammalian NAD+ homeostasis41. By using triple-quad mass spectrometry and multiple
reaction monitoring (MRM) for targeted metabolomics, we could further refine these data to
determine where mass shifts occurred, including whether Nam was labelled at the pyridine ring
or amide positions, and whether M+6 or M+7 labelling of NAD was from the NMN rather than
the adenosine phosphate moiety.
15N-Gln labelling of NAD+ biosynthesis
To test whether this scheme would lead to labelling of the NAD pool as anticipated, we first
used primary rat hepatocytes grown in vitro, to avoid contributions from the microbiome.
Hepatocytes were treated for 24 hr with 15N-glutamine (M+1) in the presence or absence of
NMN1 (M+6), or with NMN2 (M+7) (Fig. 3). Cell lysates were subject to targeted
metabolomic analysis to assess the degree of isotope incorporation into each metabolite (Fig.
219 APPENDIX
3a-e). Delivery of each of these isotopes yielded the expected M+6 and M+7 mass shifts of
NMN (Fig. 3b) as well as its de-phosphorylated counterpart NR (Fig. 3a), which is consistent
with the indirect transport of NMN26, though these data do not exclude the direct transport of
NMN via the putative transporter SLC12A842 – for this reason, the data in this investigation
could be interpreted as evidence for deamidation of NMN and/or NR, rather than NMN alone.
To test the strategy of using 15N glutamine to label NAD+ synthesis, we compared the ratio of
M+1 (nicotinamide labelled) to M+0 (endogenous) NAD+ (Fig. 3f). As expected, 15N-Gln
+ + treatment increased M+1 labelling of endogenous NAD , with M+1 NAD labelling in
untreated samples due to baseline levels of naturally occurring isotopes. High levels of M+1
NAD+ labelling (Fig. 3f) were observed in samples treated with NMN1, likely due to recycling
of the M+1 labelled Nam moiety (Fig. 3g) following the breakdown of NAD+ (Fig. 2), or by
cleavage of the NMN glycosidic bond between the ribose and nicotinamide groups. As
expected, treatment with NMN1 (M+6) and NMN2 (M+7) led to M+6 and M+7 labelling of
NAD+ (Fig. 3c). While we had hypothesised that in vivo treatment with NMN1 (M+6) and 15N-
Gln (M+1) would lead to M+7 labelled NAD+ due to the deamidation of NMN by the gut
microbiome, in these primary hepatocytes we observed that 15N-Gln co-treatment with NMN1
(M+6) increased the formation of M+7 labelled NAD+, when compared to NMN1 (M+6) alone
(Fig. 3h). In line with the expected recycling of labelled Nam from NAD+ (Fig. 2a), this
increased formation of M+7 labelled NAD+ during NMN1 (M+6) and 15N-Gln co-treatment
was matched by an identical increase in M+2 labelling of free Nam (Fig. 3i), which was re-
incorporated into the nicotinyl moiety of NAD+ (Fig. 3j). 15N-Gln treatment increased M+1
labelling at the amide position of Nam (Fig. 3i), but not the base N atom of the pyridine ring
(Nambase, Fig. 3k), which does not undergo substitution by NADS, with NMN1 (Fig. 2a)
treatment serving as a positive control for labelling at this position. Overall, these data verified
our system of labelling, and demonstrated the specificity of our targeted analytical approach,
220 APPENDIX
based on triple quadrupole mass spectrometry and MRM targeted metabolomics. As mammals
do not encode a known NMN, NR or Nam deamidase enzyme, we next sought to measure the
incorporation of labelled NMN into the NAD+ metabolome of bacteria, which can deamidate
NAD precursors including NMN36 and Nam43.
NMN deamidation by bacteria
Unlike mammals, bacteria rely on an NAD+ dependent DNA ligase that is inhibited by NMN,
the product of its own reaction37-39, resulting in the accumulation of intracellular NMN during
exponential growth44. This NMN is salvaged through the bacterial NMN deamidase PncC, yielding NaMN as a substrate for NAD+ synthesis by the Preiss-Handler pathway36. To model
whether extracellular NMN would undergo deamidation by bacteria, growth phase E. coli cultures were supplemented with NMN1 (M+6) (Fig. 2a) and subjected to targeted metabolomics of both cell lysates and extracellular culture media (Fig. 4). Consistent with the role of PncC in NMN metabolism in bacteria, treatment with labelled NMN resulted in the rapid incorporation of isotope labels into NaMN, with vastly increased labelling of NaMN compared to NMN (Fig. 4). Similarly, a role for the Nam deamidase PncA is strikingly reflected in the abundance of nicotinic acid (Na) compared to nicotinamide (Nam) in the cell pellet compared to the culture supernatant, where the ratio of Nam to Na in growth media was completely reversed. Overall, the avid uptake of NMN, followed by its rapid shunting into deamidated metabolites such as NaMN (Fig. 4) supported our hypothesis that the gut microbiome could contribute to the metabolism of orally administered NAD precursors such as NMN.
Antibiotic treatment alters NMN deamidation in vivo
To directly trace whether the increase in deamidated metabolites following NMN
administration (Fig. 1) was indeed due to the direct deamidation and incorporation of these
221 APPENDIX
metabolites, we next delivered our strategically designed isotopes into animals that had
similarly been treated with antibiotics to deplete the gut microbiome, as confirmed by
reductions in faecal DNA concentration (Supp. Fig. 1a), full-length 16S rRNA sequencing
(Supp. Fig. 1c) and reduced alpha diversity (Supp. Fig. 1e-r). Following antibiotic treatment,
animals received a single oral gavage (50 mg/kg) of the NMN1 (M+6) isotope (Fig. 2a), in
parallel with an i.p. bolus of 15N-Gln (M+1). Four hours later, animals were sacrificed and
tissues rapidly preserved for targeted metabolomic analysis (Fig. 5, Supp. Fig. 3-4). In a separate experiment, a different cohort of antibiotic treated animals (Supp. Fig. 1b, d, f-r)
received a bolus of the NMN2 (M+7) isotope (Fig. 2b) alone, following which tissues were
similarly collected 4 hr later for targeted metabolomic analysis (Fig. 6, Supp. Fig. 5, 6).
In tissues from animals treated with NMN1 (M+6), the deamidation of NMN could be
quantified by comparing the ratio of M+6 NAD+, which would assume incorporation following
the canonical route, to M+7 NAD+, which had incorporated an extra mass shift from co- treatment with 15N-Gln (M+1) (Fig. 2a, 7a). In this experiment, an increased ratio of M+7 to
M+6 labelled NAD+ would indicate the deamidation of NMN. The reason for using ratios,
rather than the overall amounts of each isotope (Fig. 5, Supp. Fig. 3, 4), is that they internally
control for differences in bioavailability within each animal. From the intact labelling of NAD+
in the GIT from NMN1 treatment, around 13% was M+7 labelled (Fig. 7b, c). Importantly,
these data likely underestimates incorporation via the deamidated route, as this scheme relied
on the availability of exogenous 15N-Gln relative to the endogenous pool of unlabelled Gln,
which composed only 9-13% of the total plasma Gln pool at the 4 hr timepoint (Fig. 7k).
Consistent with our hypothesis, the ratio of M+7 to M+6 labelling in the GIT was reduced in
antibiotic treated animals (Fig. 7b), suggesting reduced deamidation of orally administered
NMN when contributions from the microbiome were reduced. This was reflected by a
reduction in the ratio of M+2 to M+1base labelled Nam in the GIT, liver and plasma (Fig. 7d-f),
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however this change also likely reflected reduced contributions from the bacterial nicotinamide
de-amidase PncA43 following antibiotic treatment41. This change in Nam labelling carried into
the M+2 to M+1 ratio of NMN (Fig. 7g, h) and NAD+ (Fig. 7i, j), with reduced labelling ratios
of these recycled isotopes during antibiotic treatment likely reflecting a combination of
possible NMN/NR deamidation (Fig. 7a, b), and contributions from the bacterial Nam
deamidase PncA41.
To complement this approach, in the NMN2 (M+7) experiment (Fig. 2b), we would anticipate
that deamidation by the microbiome would result in loss of the 15N amide label, resulting in
the formation of M+6 NAD+ at the expense of M+7 NAD+ (Fig. 2b, 7l). In contrast to the
previous NMN1 experiment, the ratio of M+7 to M+6 NAD+ would instead decrease as the
rate of deamidation increased. Further, interpretation of deamidation in this NMN2 experiment
was not limited by the availability of exogenous 15N-Gln relative to a large, endogenous pool
of Gln, as was the case with the NMN1 experiment (Fig. 7a, k). In this experiment, the ratio of
M+7 to M+6 NAD+ was around 3:1 (Fig. 7m, n), suggesting that around 25% of orally
administered NMN undergoes deamidation prior to its intact incorporation into NAD+. In
agreement with the previous experiment, the ratio of M+7 to M+6 labelled NAD+ was increased in the GIT and liver of antibiotic treated animals (Fig. 7m, n). This was similarly matched by an increased ratio of M+2 to M+1base labelled Nam in the GIT (Fig. 7o), liver (Fig. 7p) and plasma (Supp. Fig. 7q) of antibiotic treated animals, reflecting decreased incorporation following deamidation, though this could instead be due the deamidation of Nam rather than
NMN. As in the previous experiment, these labels were recycled into M+2 labelled NMN (Fig.
7r, s) and NAD+ (Fig. 7t, u). In addition to differences in the isotope labelling of NAD+ (Fig.
7), these experiments replicated the inverse relationship between NaMN and NMN levels
following antibiotic treatment (Fig. 5b, e, Fig. 6b, e) observed in our earlier experiment with
unlabelled NMN (Fig. 1b, h, n). Overall, these data from two complementary isotope labelling
223 APPENDIX
approaches support the concept that orally delivered NMN or NR can undergo deamidation
prior to incorporation, and a role for the microbiome in mediating this. While these data could
in part explain the spike in the de-amidated metabolites NaMN and NaAD following treatment
with the amidated precursors NR25 or NMN (Fig. 1), it is important to note that when measured
as a proportion of the overall NAD+ pool, the contribution of both M+7 and M+6 intact labelled
NAD+ was small. Partially labelled NAD+ (M+2) was around 10-fold more abundant than intact
labelled NAD+ (M+7) (Fig. 5d, 6d, Supp. Fig. 3-6), indicating either cleavage of the labile glycosidic bond of NMN, or rapid recycling of NAD+45. Following cleavage of the glycosidic
bond to release free Nam, its deamidation in the GIT46,47 by the bacterial enzyme PncA41 also
likely contributes to these changes.
Exogenous NMN boosts the endogenous NAD+ metabolome
The abundance of partially labelled NAD+ with labelling at the Nam position only is consistent
with previous findings45 that orally delivered NMN and NR undergo cleavage at the glycosidic bond to release free Nam, with only a small proportion of orally delivered material being incorporated into tissues intact. Given that NMN and NR are overwhelmingly incorporated in the form of free Nam45, why do the downstream biological effects of NR or NMN differ from
the delivery of Nam25,48-50, a widely available nutrient present in dietary sources? One basic
assumption is that exogenous NAD precursors increase levels of NAD+ and other metabolites
due to their direct incorporation, as expected by classic mass-balance models. Following the
delivery of near 100% isotope labelled material, we found that the increase in metabolites such
as NR includes the increased production of endogenous, unlabelled metabolites. For example,
treatment with NMN1 or NMN2 in antibiotics treated animals increased levels of unlabelled
NaR (Fig. 5f, 6f, Supp. Fig. 3f, 5f) in the GIT, and increased unlabelled NR (Fig. 5b, 5p, 6b,
6p Supp. Fig. 3b, 5b) and NaAD (Fig. 5n, 6n, Supp. Fig. 4g, 6g) in the liver. Notably, treatment
with labelled NMN in antibiotics treated animals increased the production of unlabelled NR in
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the GIT by over 3-fold, where this single metabolite accounted for the vast majority of the
entire NAD metabolome in the gut, as summarised in Figure 5p and 6p.
Host-microbe interactions in the bioavailability of orally delivered NAD+ precursors
Another unexpected aspect of these data was the overall increase in levels of these metabolites as a result of antibiotics treatment alone, which more than doubled the labelling of the metabolites NMN, NR, NAD+ and Nam (Fig. 1g-i, 5a-d, 6a-d, Supp. Fig. 3h-k, 4h-k, 5h-k, 6h- k). This increase even occurred in unlabelled metabolites in animals that did not receive exogenous NMN (Supp. Fig. 3a-d, 5a-d). When NMN1 (M+6) was delivered, the incorporation of exogenous labels into NAD+ metabolites was vastly increased in antibiotic treated animals,
in the case of NR in the gut, by an order of magnitude (Fig. 5c, 5p, Supp. Fig. 3i), a trend that
was recapitulated in a separate cohort of animals receiving the NMN2 (M+7) isotope (Fig. 6b,
6p, Supp. Fig. 5i) and in animals that received unlabelled NMN (Fig. 1g). The overwhelming
abundance of NR as the dominant NAD metabolite in the GIT, especially following NMN
delivery in antibiotics treated animals is worthy of later investigation, as the abundance of this
single metabolite was greater than all other NAD+ metabolites combined, including NAD+
itself (summarised in Fig. 5p, 6p). Together, the striking increase in the uptake and overall
abundance of both labelled and unlabelled NAD metabolites in antibiotics treated animals
suggests that the microbiome could be in competition with mammalian tissue for the uptake of
orally administered, exogenous NAD precursors, and the uptake of NAD precursors from
dietary sources. Future studies should measure isotope labelling of NAD+ metabolites in faecal
contents of mice to confirm whether these compounds are being utilised by the microbiome,
rather than being excreted via other mechanisms, and should use animals in which the
microbiome has been reconstituted to control for the effects of antibiotics treatment.
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Evidence for NMN uptake following dephosphorylation into NR
NMN uptake can occur following the dephosphorylation of NMN into NR by the cell surface
enzyme CD73, prior to uptake by ENT nucleoside transporters and re-phosphorylation into
NMN inside the cell by NRK1/2 (Fig. 8)26,27. Alternatively, the solute carrier protein SLC12A8
has been described as a dedicated NMN transporter42. As with CD73 and ENT, SLC12A8 is
located on the apical side of the intestinal tissue. As both mechanisms could co-exist, the
question is the degree to which each mechanism contributes to the uptake of NMN51. If the direct route via SLC12A8 prevailed, we would expect to see high levels of labelled (M+7 or
M+6) NMN, with lesser uptake of labelled NR. In contrast, if the indirect transport of NMN following its dephosphorylation into NR was dominant, there would be a higher levels of NR labelling. In primary hepatocytes (Fig. 3), NMN1 (M+6) treatment resulted in near complete labelling of the NR pool (Fig. 2b), with slightly lesser labelling of the NMN pool (Fig. 2a).
Strikingly, in vivo treatment showed strong labelling of the NR, but not NMN pools (Fig. 5b, c; Fig. 6b, c; Supp. Fig. 3a-b, h-i; Supp. Fig. 5a-b, h-i; Supp. Fig. 7). Intact (M+6 or M+7) labelled NR levels (Supp. Fig. 3i, 5i) were five-fold higher than endogenous NR (M+0) (Fig.
5c, 6c, Supp. Fig. 3h, 5h; Fig. 8), which increased to a ten-fold greater enrichment with
antibiotics treatment (Supp. Fig. 8). In stark contrast, only around 5% of the NMN pool was
M+7 labelled (Fig. 5b, Fig. 6b; Supp. Fig. 3a, h; Supp. Fig. 5a, h, Fig. 8). The inability of
exogenous NMN to displace the endogenous NMN pool, combined with the surge of labelled
NR, suggests that NMN uptake bypasses direct transport, and would instead support the
dephosphorylation of NMN into NR to facilitate its intestinal absorption (Fig. 8). If direct
transport of NMN does occur, it (along with NR) is in competition with the microbiome, as
even when M+6 or M+7 labelling of NMN was observed at low levels, this only occurred in
antibiotic treated animals (Fig. 5b, 6b). An important caveat of this interpretation is that limited
availability of isotope labelled material meant that this study used a single time point, rather
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than a time course which also encompassed very early timepoints, possibly missing the minute-
order kinetics of direct NMN transport that were previously reported20,52.
Chronic NMN treatment does not alter microbial diversity
Given the potential role for host-microbiome interactions in the metabolism of orally delivered
NMN, it might be expected that NMN treatment would alter the composition of the
microbiome. To test this, we treated aged, 97-week old animals with chronic NMN dosing via
addition to drinking water (~400 mg/kg/day) for 8-10 weeks prior to cull7,9,11,53. 16S rRNA
long-read sequencing revealed no overall change in the alpha diversity of the gut microbiome
(Fig. 9), however there were a number of changes among individual species, available as a
Supplementary File.
DISCUSSION
Together, this work provides evidence for the partial incorporation of exogenous NMN into the NAD metabolome via the deamidated route, and for contributions of the gut microbiome to the metabolism of exogenous NMN. This is in line with recent findings around the role of
Nam deamidation by bacteria41, however a role for the microbiome in the uptake of exogenous
NAD precursors has not been described. It will be interesting to determine whether this relationship persists for other precursors, or is unique to NMN. Rather than being evidence for a “competition” relationship, differences in the uptake of exogenous NMN could reflect its role as an inhibitor of bacterial DNA ligase37-39, and bacterial mechanisms to prevent its accumulation. In addition to NMN deamidase enzymes, this could include a role for bacterial
NAD glycohydrolase enzymes, and/or bacterial SARM-like enzymes54.
While intravenous delivery of NR or NMN results in a small degree of intact assimilation into peripheral tissues such as the liver, kidney and muscle, oral delivery of NMN results in hepatic cleavage at the glycosidic bond yielding free nicotinamide due to the action of the liver 45. Our
227 APPENDIX results were in close alignment with those findings45, where the ratio of intact M+7 or M+6 to
M+0 unlabelled NAD+ was around 2%, whereas the ratio of M+2 labelled to M+0 unlabelled
NAD+, presumably as a result of incorporation of free Nam, was over 10% (Fig. 5d, k; 6d, k).
Given this evidence for the decomposition of NMN into free Nam prior to its uptake, a key question is why downstream precursors in NAD+ synthesis such as NMN and NR lead to different outcomes compared to Nam alone48-50. A surprising aspect of these results was that treatment labelled NMN led to an increase in unlabelled NAD metabolites. In the case of both
NR and NaR, treatment with NMN1 (M+6) or NMN2 (M+7) led to a stark increase in endogenous (M+0) levels, particularly in antibiotics treated animals (Fig. 5c, f, 6c, f, Supp. Fig.
3b, f, 5b, f). We also observed that exogenous NMN increased liver NaAD levels (Fig. 1f, 5g, n, 6g, n), as was previously reported for exogenous NR treatment25, however the majority of this increase was from unlabelled NaAD (Supp. Fig. 4g, 6g). These results were similar to those of Trammel et al25, where isotope tracing of double-labelled NR showed that while total NaAD levels increased by over 40-fold following NR treatment, only around 45% of this NaAD was isotope labelled – with the endogenous origin of the remaining 55% remaining unexplained.
We argue that these findings run against the assumed model that exogenous NAD+ precursors raise NAD+ levels through their direct incorporation into the NAD metabolome, and instead could suggest that treatment with exogenous precursors could indirectly trigger endogenous
NAD+ biosynthesis. The mechanism for this is not yet clear, though given the profound effect of antibiotic treatment, in particular for the overwhelming abundance of NR in the gut (Fig. 5c, p, 6c, p, Supp. Fig. 3b, 5b), it is likely to involve interplay with the gut microbiome. One possibility for the changes in endogenous NAD+ metabolites following exogenous NMN/NR treatment could be that exogenous NMN or NR treatment triggers unknown signalling pathways that indirectly alter endogenous NAD metabolism, rather than the direct incorporation of exogenous material.
228 APPENDIX
Another explanation is that increased substrate levels alter the in vivo kinetics of NAD
biosynthetic enzymes, increasing the utilisation of endogenous substrates. An important
question regarding NR in particular is how its endogenous production is increased by
exogenous NMN. NR is available from dietary sources27, and is an intermediate in the uptake
of extracellular NMN55,56. NMN accumulation in neurons can trigger cell death through the
NADase SARM157, and disposal of NMN through its adenylation into NAD+ can protect
against neuronal death58. It is possible that exogenous NMN triggers pathways that degrade
endogenous NMN into NR, which could act as a reservoir for NAD precursors, however CD73,
the NMN ectonucleotidase that carries this out, sits on the extracellular face of the plasma
membrane55,56 rather than the cytosol. In addition, the sheer molar quantity of unlabelled NR that we observed in the gut (Fig. 1g, 5c, 6c) relative to other metabolites (Fig. 5p, 6p) challenges this idea. NAD homeostasis is tightly maintained within a defined range59, and the activity of
NMNAT enzymes that carry out the last step of NAD biosynthesis is reversible60. It is possible
that exogenous NAD precursors push the equilibrium of this step in the opposite direction,
increasing endogenous NMN production from NAD+, though how intracellular NMN could be
dephosphorylated to an NaR/NR reservoir in mammals is unknown. This concept of increased
NAD breakdown during treatment with exogenous NMN in young animals is also supported
by the increased formation of unlabelled Nam in plasma (Fig. 5o, 6o). Further, while labelled
NMN treatment in vitro (Fig. 3) results in the formation of intact labelled NAD+, this occurs at
the cost of unlabelled NAD+, for a net zero change in total NAD levels (Fig. 3c), suggesting
that when NAD+ is replete, the utilisation of exogenous NMN into newly synthesised NAD+
results in a commensurate breakdown of existing material – potentially explaining the
formation of unlabelled metabolites such as NR during treatment with exogenous, labelled
NMN.
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Rather than degrading or recycling existing metabolites into NR, another possibility is that
exogenous NMN or NR could trigger a currently unknown step in mammals that leads to
endogenous NR production. While NR can be produced by the reversible phosphorolysis of
Nam and ribose-5-phosphate by a purine nucleophosphorylase (PNP) in E. coli61, this step is
irreversible in mammals56,62,63, and other potential steps involved in endogenous NR
production in mammals are unknown. Further work is needed to understand how this occurs,
for example, whether it is due to the acute up-regulation of NAD+ biosynthetic enzymes, the
time-scale by which this increase occurs, and a direct comparison of different isotope labelled
NAD+ precursors to identify which metabolites trigger the production of endogenous
metabolites under normal circumstances and during depletion of the microbiome. Regardless,
the ability to trigger the production of endogenous NAD metabolites could explain why
exogenous NR and NMN treatment lead to differences in pharmacokinetics, metabolite
production and therapeutic effects when compared to Nam alone48-50, despite their rapid
metabolism into free Nam by the liver45 (Fig. 4, 5).
Another speculative idea is the existence of a signalling pathway in the GIT that is sensitive to
both exogenous NAD precursors and to microbial metabolites, which can mediate endogenous
NAD metabolism. Metabolite sensing members of the G-protein coupled receptor (GPCR) family are putative candidates for this role as there are already GPCRs known to respond to extracellular nicotinic acid and to NAD+ itself64. One possible candidate is GPR109a, which
acts as a receptor both for nicotinic acid65 and for butyrate, released from the microbial
fermentation of dietary fibre66. This could link the observations including the deamidation of
orally derived NAD precursors into their acid equivalents, a role for microbiome depletion in
triggering the production of endogenous NAD metabolites, and evidence for the poor
incorporation of intact NR or NMN into the NAD metabolome. Identifying cell surface
230 APPENDIX receptors and downstream signalling pathways in the gut that are sensitive to both microbial metabolites and exogenous NAD+ precursors will be a key goal of testing our hypothesis around the role of exogenous metabolites in endogenous NAD+ metabolism. Together, these findings regarding the deamidation of NMN and its effects on the endogenous NAD metabolome have profound importance for the therapeutic development of NAD precursors.
231 APPENDIX
Methods
Methods are available in Supplementary Material, with raw data available on our Mendeley data site.
Acknowledgements
Funding was from the National Health and Medical Research (NHMRC) of Australia as a
Career Development Fellowship APP1122484 to LEW, and sponsored research from Jumpstart
Fertility. We wish to thank anonymous donors for philanthropic support.
Author contributions
LJK conducted experiments, analysed data, prepared figures, wrote manuscript. TJC conducted microbiome analyses. EWKP, TTC, JW prepared isotope labelled NMN. SPT and DAS provided critical feedback and interpretation. LEQ conducted experiments, extracted and analysed data, wrote manuscript. LEW conceived of and designed study, obtained funding, supervised experiments, analysed data, prepared figures, wrote manuscript.
Declaration of interests
EWKP and JW are employees and shareholders of GeneHarbor Biotechnologies. SPT is the
CEO of Jumpstart Fertility, which is developing NAD+ raising compounds for therapeutic use.
LEW and DAS are co-founders, shareholders, directors and advisors of Jumpstart Fertility and the Life Biosciences group which includes Jumpstart Fertility, Continuum Biosciences,
Senolytic Therapeutics, Selphagy, and Animal Biosciences. LEW and DAS are also advisors to and shareholders in the EdenRoc group of companies, which includes Metro Biotech NSW and Metro International Biotech, Arc-Bio, Dovetail Genomics, Claret, Revere Biosciences, and
Liberty Biosecurity. LEW is an advisor and shareholder in Intravital Pty Ltd. DAS is an
232 APPENDIX inventor on a patent application that has been licensed to Elysium Health. Updated affiliation are at https://genetics.med.harvard.edu/sinclair-test/people/sinclair-other.php.
233 APPENDIX
Figure 1
Figure 1. NMN treatment leads to the microbiome-dependent formation of deamidated
NAD+ metabolites in vivo. Mice treated with antibiotics (Abx) to ablate the gut microbiome
were administered a single dose of unlabelled nicotinamide mononucleotide (NMN) (500
mg/kg, oral gavage). Gastrointestinal tissue (GIT) (a-c) and liver tissue (d-f) were subject to targeted mass spectrometry to quantify the deamidated metabolites (a-f) nicotinic acid riboside
(NaR) (a, d), nicotinic acid mononucleotide (NaMN) (b, e) and nicotinic acid adenine
dinucleotide (NAAD) (c, f), as well as their amidated counterparts (g-l) nicotinamide riboside
234 APPENDIX
(NR) (g, j), NMN (h, k) and nicotinamide adenine dinucleotide (NAD+) (i, l). These data were
then expressed as ratios between de-amidated and amidated counterparts in GIT (m-o) and liver
(p-r). Data analysed by 2-way ANOVA with Sidak’s post-hoc test, exact p-values and F values in supplementary files. N=4-5 animals per group, *p<0.05, **p<0.01, ***p<0.001,
****p<0.0001.
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Figure 2
Figure 2. Scheme for assimilation of isotope labelled NMN into the NAD+ metabolome.
NMN (box on left) was labelled with 13C at all 5 carbon positions of the ribose moiety
(highlighted in red) and with 15N at the base (highlighted in blue) and amine (highlighted in orange) of the nicotinamide moiety for a total mass shift of M+7. The expected mass shift for
236 APPENDIX each species is shown, with steps catalyzed by mammalian enzymes in solid black lines, and steps that are only known to be carried out by bacterial enzymes shown with dashed lines.
237 APPENDIX
Figure 3
Figure 3. 15N-glutamine labelling of NAD+ synthesis. Primary hepatocytes were treated with
NMN1 (M+6) or NMN2 (M+7) isotopes (200 µM) in the presence of unlabelled or amide labelled 15N-glutamine (M+1) (4 mM) for 24 hr, to measure the degree of NAD synthesis and
the incorporation of exogenous NMN via the de novo pathway, which incorporates the amide
238 APPENDIX
label of 15N-Gln through the enzyme NAD synthetase (NADS). Exogenous NMN1 and NMN2
isotopes in the presence of 15N-Gln led to the expected isotopic labelling of a) NR, b) NMN, c)
+ NAD d) Nam and e) Gln. Ratios of f) M+1amide labelled to unlabelled NAD, g) M+7 labelled
15 NAD+, h) M+2 labelled NAD+ and i) M+1amide Nam were consistent with labelling by N-
Gln, with no change in j) M+1base labelled Nam acting as a negative control, as no change in
labelling at the ring position is expected. This resulted in l) M+2 labelled Nam during NMN1
and 15N-Gln treatment. Data analysed by 2-way ANOVA with Sidak’s post-hoc test, n=3 biological replicates.
239 APPENDIX
Figure 4
Figure 4. NMN deamidation in bacteria. Liquid cultures of E. coli OP50 bacteria were supplemented with M+6 labelled NMN1 (0.1 mM) at inoculation of a fresh culture. Samples were taken at time 0 (after NMN), 140, 160 and 180 minutes after NMN supplementation.
240 APPENDIX
Following separation of the culture supernatant (top) from the cell lysate (bottom), metabolites
were extracted and subjected to targeted LC-MS/MS mass spectrometry to detect the incorporation of the M+6 isotope label into NMN, NR, NaMN and NAD+ as well as M+1
labelling of nicotinamide (Nam) and nicotinic acid (Na) in both the culture supernatant (top)
and cell lysates (bottom). Data represents mean ± s.d. (n=3-5 samples per time point).
241 APPENDIX
Figure 5
Figure 5. Incorporation of NMN1 (M+6) isotope into the NAD metabolome in vivo.
Animals were treated with antibiotics (Abx) to deplete the microbiome, followed by an oral gavage (50 mg/kg) of NMN1 (M+6) with adjacent i.p. administration of 15N-Gln (735 mg/kg,
10ml/kg body weight). Four hr later, GIT (a-g), liver (h-n) and plasma (o) were rapidly
preserved for targeted metabolomics analysis to identify labelling of (a, h, o) Nam, (b, i) NMN,
(c, j) NR, (d, k) NAD+, (e, l) NaMN, (f, m) NaR and (g, n) NaAD. Data presented as stacked
bars of each isotopologue, with raw data points for each isotopologue overlaid on bar charts.
242 APPENDIX
(p, q) Relative molar abundance of all NAD metabolites in (p) GIT and (q) liver, including
endogenous (M+0), intact labelled and partially labelled (“part”) isotopologues. n=3-4 animals per group, n.d. = not detected.
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Figure 6
Figure 6. Incorporation of NMN2 (M+7) into the NAD metabolome in vivo. Animals were treated with antibiotics (Abx) to deplete the microbiome, followed by an oral gavage (50 mg/kg) of NMN2 (M+7). Four hr later, GIT (a-g), liver (h-n) and plasma (o) were rapidly preserved for targeted metabolomics analysis to identify labelling of (a, h, o) Nam, (b, i) NMN,
(c, j) NR, (d, k) NAD+, (e, l) NaMN, (f, m) NaR and (g, n) NaAD. Data presented as stacked bars of each isotopologue, with raw data points for each isotopologue overlaid on bar charts.
(p, q) Relative molar abundance of all NAD metabolites in (p) GIT and (q) liver, including
244 APPENDIX endogenous (M+0), intact labelled and partially labelled (“part”) isotopologues. n=3-4 animals per group n.d. = not detected.
245 APPENDIX
Figure 7
Figure 7. Contribution of the microbiome to NMN deamidation. Antibiotic treated animals were orally administered with the a) NMN1 (M+6) isotope in the presence or absence of 15N- glutamine (M+1), with the formation of M+7 labelled NAD+ or M+2 labelled Nam reflective
246 APPENDIX
of incorporation following deamidation and reamidation by the enzyme NADS. Antibiotic
treatment reduced (b, c) M+7 labelling of NAD+, (d-f) M+2 labelling of nicotinamide (Nam),
(g, h) M+2 labelling of recycled NMN, and (i-j) M+2 labelling of recycled NAD+, which
reflects the incorporation of recycled Nam. Data are expressed as ratios to M+6 or M+1 NAD+
and M+1base Nam, which are the expected isotope products of NMN or NR assimilation via the canonical (amidated) route. Incorporation of this extra label was limited by the availability of
15N-glutamine (M+1) as k) a proportion of the endogenous glutamine pool in plasma. In a separate cohort, animals were treated with l) NMN2 (M+7), where loss of the amide 15N label
+ to form M+6 labelled NAD or M+1base labelled Nam would reflect deamidation. Antibiotic treatment protected M+7 NAD+ (m, n), M+2 Nam (o-q), and M+2 NAD+ (n, o) against loss of
the NMN2 15N amide compared to untreated animals, which carried into M+2 labelling of recycled NMN, and (i-j) M+2 labelling of recycled NAD+. Tissues shown in the left column
(b, d, g, I, m, o, r, t) are gastrointestinal tract (GIT), middle column (c, e, h, j, n, p, s, u) are liver, right column (f, k, q) are plasma. NMN1 and NMN2 experiments were run in separate cohorts of animals, each measurement represents tissue from a separate animal. Comparisons of isotope treated groups with or without antibiotic treatment were analysed by Mann-Whitney
U-test, *p=0.0286, n=4 animals per group.
247 APPENDIX
Supplementary Figure 8
Supplementary Figure 8. Isotope labelled NMN treatment results in greater labelling of the NR than NMN pool, suggesting indirect uptake. (a) The two proposed mechanisms for
NMN uptake are either directly through the putative NMN transporter SLC12A8, or indirectly by dephosphorylation into NR via the ecto-5’-nucleotidase CD73 which is present on the apical side of intestinal cells. To compare the contributions of either direct or indirect transport mechanisms, the contribution of isotope labelled NMN to the overall pool of each metabolite is shown for the intestinal tissue of the NMN2 cohort (data from Fig. 6). Error bars are s.d., each data point represents tissue from a separate animal.
248 APPENDIX
Figure 9
Figure 9. Effects of chronic NMN treatment on the aged gut microbiome. Aged (97-week old) male and female mice were treated with NMN through addition to drinking water (~400 mg/kg/day) for 8-10 weeks prior to cull, and faecal microbiome samples subject to long-read
16S rRNA sequencing. (a) Stacked bar plots represent the total reads and relative abundance of bacterial taxa at the genus level for untreated or NMN treated animals, also shown as differential expression by the (b) genus level and (c) by operational taxonomic unit (OTU) level. There was no change in microbiome diversity, shown by the (d) Shannon alpha diversity index.
249 APPENDIX
SUPPLEMENTARY FIGURES
250 APPENDIX
Supplementary Figure 1
Supplementary Figure 1. Antibiotics treatment ablates the gut microbiome. Following
antibiotic treatment in the NMN1 and NMN2 mouse cohorts, (a-b) DNA was extracted from
faeces to measure changes in DNA concentration. Uniform amounts of DNA were then
subject to (c-d) full-length 16S rRNA Nanopore sequencing, with species abundance shown
here at the genus level. Sequencing revealed a reduction in (e-f) operational taxonomic units
(OTUs), the (g-h) Chao1 and (i-j) ACE species richness indices, and the (k-l) Shannon, (m-n)
Simpson, (o-p) Inverse Simpson and (q-r) Fisher diversity indices. Data shown are non-
251 APPENDIX rarefied; rarefication showed identical results (data not shown). Each data point represents samples from a separate animal
252 APPENDIX
Supplementary Figure 2
Supplementary Figure 2. Chromatogram of isotope labelled NMN and glutamine (Gln) using MRM LC-MS/MS. The above chromatograms represent individual peaks (ion count) for a) 100µM unlabelled NMN in combined NAD metabolite standard curve mixture, b) M+6 labelled NMN1 and c) M+7 labelled NMN2, as well as d) unlabelled and 15N-amide labelled glutamine.
253 APPENDIX
Supplementary Figure 3
254 APPENDIX
Supplementary Figure 3. Incorporation of NMN1 (M+6) into the GIT (raw data for Fig.
5). As in Figure 4 of main text, animals were treated with antibiotics (Abx) and M+6 isotope
labelled NMN1 followed by metabolomics analysis of intestinal tissue as described in Figure
1 of main text. Animals were also intraperitoneally injected with a concurrent bolus of 15N- amide labelled glutamine (735 mg/kg, 10ml/kg body weight). The left column (a-g) represents the abundance of unlabelled, endogenous species, middle column (h-n) represents intact label incorporation, right column (o-u) represents partial labelled species from the recycling of metabolites, as predicted in Fig. 2a. Data analysed by 2-way ANOVA with Sidak’s post-hoc test. n=3-4 animals per group, each data point represents a separate animal, error bars are SD.
255 APPENDIX
Supplementary Figure 4
256 APPENDIX
Supplementary Figure 4. Incorporation of NMN1 (M+6) into the liver (raw data for Fig.
5). As in Figure 4 of main text, animals were treated with antibiotics (Abx) and M+6 isotope
labelled NMN1 followed by metabolomics analysis of intestinal tissue as described in Figure
1 of main text. Animals were also intraperitoneally injected with a concurrent bolus of 15N-
amide labelled glutamine (735 mg/kg, 10ml/kg body weight). The left column (a-g)
represents the abundance of unlabelled, endogenous species, middle column (h-n) represents
intact label incorporation, right column (o-u) represents partial labelled species from the
recycling of metabolites, as predicted in Fig. 2a. Data analysed by 2-way ANOVA with
Sidak’s post-hoc test. n=3-4 animals per group, each data point represents a separate animal, error bars are SD.
257 APPENDIX
Supplementary Figure 5
258 APPENDIX
Supplementary Figure 5. Incorporation of NMN2 (M+7) into the GIT (raw data for Fig.
6). Animals were treated with antibiotics (Abx) and M+7 isotope labelled NMN2 followed by
metabolomics analysis of GIT tissue as described in Figure 6 of main text. Each data point
represents measurements from a different animal. The left column (a-d) represents the
abundance of unlabelled, endogenous species, middle column (e-h) represents intact label incorporation, right column (i-l) represents partial labelled species from the recycling of metabolites, as predicted in Fig. 1. Metabolites assayed are nicotinamide riboside (NR), nicotinamide adenine dinucleotide (NAD+), nicotinamide (Nam) and nicotinamide
mononucleotide (NMN). Data are analysed by two-way ANOVA with Sidak’s multiple
comparisons test. Data are mean ± s.d. (n=3-5 mice per group). *p<0.05, **p<0.01,
***p<0.001, ****p<0.0001, ns=not significant.
259 APPENDIX
Supplementary Figure 6
260 APPENDIX
Supplementary Figure 6. Incorporation of NMN2 (M+7) into the liver (raw data for Fig.
6). Animals were treated with antibiotics (Abx) and M+7 isotope labelled NMN2 followed by
metabolomics analysis of liver tissue as described in Figure 6 of main text. Each data point
represents measurements from a different animal. The left column (a-d) represents the
abundance of unlabelled, endogenous species, middle column (e-h) represents intact label incorporation, right column (i-l) represents partial labelled species from the recycling of metabolites, as predicted in Fig. 1. Metabolites assayed are nicotinamide riboside (NR), nicotinamide adenine dinucleotide (NAD+), nicotinamide (Nam) and nicotinamide
mononucleotide (NMN). Data are analysed by two-way ANOVA with Sidak’s multiple comparisons test. Data are mean ± s.d. (n=3-5 mice per group). *p<0.05, **p<0.01,
***p<0.001, ****p<0.0001, ns=not significant.
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SUPPLEMENTARY METHODS
262 APPENDIX
Synthesis of isotope labelled NMN
The isotopes used here were generated through a two-step process starting with the custom
synthesis of nicotinamide labelled with 15N at the nitrogen base and amide positions. This
custom isotope labelled version of nicotinamide was then used with [U5]-13C- ribose which
was 13C labelled at all five carbon positions (Cambridge Isotope Laboratories, cat. no. CLM-
3652) and ATP in an enzyme-based protocol using recombinant phosphoribosyl synthetase
(PRS) and recombinant nicotinamide phosphoribosyl transferase (NAMPT) to synthesise
NMN. The two enzymes were added into the reaction buffer that contains 1 mM ribose, 1 mM
nicotinamide, 3 mM ATP, 1 mM dithiothreitol, 10 mM MgCl2 and 50 mM Tris-HCl (pH 7.5)
and incubated at 37℃ for 30 min. The reaction was terminated with the addition of 0.01%
Trichloroacetic acid (TCA). The purification was proceeded with size-exclusion columns and
ion exchange columns. Isotope labelled NMN samples of >95% purity were concentrated by
lyophilization, and labelling confirmed by mass spectrometry (Supp. Fig. 2).
Animal experiments
All experiments were performed according to procedures approved by UNSW Animal Care
and Ethics Committee (ACEC) under ethics protocol 18/134A. The UNSW ACEC operates
under the animal ethics guidelines from the National health and Medical Research Council
(NHMRC) of Australia. Mice were fed standard chow ad libitum and housed under a 12-hr
light/12-hr dark cycle in a temperature-controlled room (22 ± 1 °C) at 80% humidity in
individually ventilated cages. Four-week old female C57BL/6J mice were acclimatised for one
week prior to treatment and body weight matched before random assignment into groups. For
antibiotic treatment, mice were administered a cocktail of antibiotics consisting of vancomycin
(0.5 g/L; Sigma SBR00001), neomycin (1 g/L; Sigma N6386), ampicillin (1 g/L; Sigma
A9393) and metronidazole (1 g/L; Sigma, M3761) (VNAM) with addition of sucrose (3 g/L;
263 APPENDIX
Bundaberg Sugar) to increase palatability for 4 days, and switched to ampicillin (1 g/L) with sucrose (3 g/L) for an additional week, which can reduce gut bacterial density by 1000-fold 67.
During treatment with the VNAM combination there was a reduction in water consumption
(below), which was the reason for the subsequent switch to ampicillin alone. Sucrose treatment
(3 g/L) was maintained as a vehicle control in animals that did not receive antibiotic treatment.
Above: Water intake and body weight of mice during antibiotics treatment.
To maintain consistency between the three cohorts presented here (unlabelled NMN – Fig. 1,
NMN1 – Fig. 5, NMN2 – Fig. 6), this antibiotic treatment protocol was used for all in vivo
experiments (Figs. 1, 5, 6). For NMN treatment, mice received a single oral gavage of NMN1
(Fig. 5) or NMN2 (Fig. 6) isotopes at 50 mg/kg, or for unlabelled NMN (Fig. 1) at 500 mg/kg,
with water vehicle used as a control. Four hours later, animals were placed under anaesthesia,
and blood was obtained by cardiac puncture, followed by euthanasia by cervical dislocation,
rapid dissection and snap freezing of tissues. Gavages were staggered between mice in
alternating treatment groups to avoid any experimental bias. On the day of cull mice were all
5-6 weeks old. Differences in NMN dosing between unlabelled and isotope labelled NMN were
due to limited availability of isotope labelled NMN.
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Blood plasma collection and preparation for mass spectrometry
Approximately 1 mL of blood was collected via cardiac puncture in anaesthetised mice (1-2%
isoflurane) into 1.5 mL eppendorf tubes prefilled with 10 µL of EDTA (0.5 M) and mixed
thoroughly with a pipette to prevent clotting. Blood samples were then spun at 2000 g for 10
min and the top layer was transferred to a new tube and snap frozen immediately in liquid
nitrogen. All samples were stored in -80°C until further processing. On the day of sample
acquisition plasma samples were thawed on ice and 20 µL of plasma was added to 80 µL of
extraction buffer (acetonitrile:methanol) with an internal standard mixture containing MES,
CSA and thymine-d4. Samples were vortexed and centrifuged at 16,000g for 10 mins at 4°C
and the supernatant was transferred to a new eppendorf tube and dried down completely using
a speed vacuum concentrator (Savant SpeedVac ® SPD140DDA, Thermo Scientific). The
resulting pellet was then resuspended in 30 µL of LC-MS-grade water and centrifuged as above and the supernatant analysed promptly by LC-MS.
Gastrointestinal and liver tissue collection and preparation for mass spectrometry
Intestinal contents (small intestine and colon without cecum) were resected and flushed with
ice cold 1 x phosphate buffered saline (PBS) to clear faecal contents before snap freezing
immediately with liquid nitrogen. Livers were resected and weighed before being rinsed in ice
cold 1x PBS and snap frozen in liquid nitrogen. All tissue samples were stored in -80 °C until
further processing. Frozen tissue samples were crushed using a mortar and pestle on liquid
nitrogen and approximately 50 mg was weighed into tubes containing ceramic beads (Precellys,
Bertin Technologies, France) to which 500 µl of cold (-30 °C) extraction buffer
(acetonitrile:methanol:water, 2:2:1) with internal standard mixture as above was added. All samples were homogenised using an automated tissue homogeniser (Precellys24, Bertin
Technologies, France) at 5,000-6,000 rpm for 15 seconds and immediately centrifuged at
265 APPENDIX
16,000g for 10 mins at 4 °C. The supernatant was transferred to a new tube and dried down completely using a speed vacuum concentrator (Savant SpeedVac ® SPD140DDA, Thermo
Scientific). All samples were resuspended in 50 µL of LC-MS-grade water, centrifuged as above and the supernatant analysed promptly by LC-MS.
Bacterial culture and NMN treatment A stab culture of the E. coli strain OP50 was inoculated into sterile Luria-Bertani (LB) broth
(10 g/L tryptone, 5 g/L yeast and 10 g/L sodium chloride in deionized water) under aseptic conditions and incubated overnight at 37℃ on a shaking platform set at 200 rpm. To measure the growth rate of E. coli, the overnight culture was sub-cultured (1:200) into sterile LB broth in a new flask and the optical density was measured at 600 nm (OD600) every 20 minutes
(approximate doubling time) and samples were collected during the early-mid exponential growth phase (OD600 < 0.70), as bacterial enzymes are more active during exponential growth phase than stationary phase 68. For samples, the overnight culture was sub-cultured (1:200) and aliquoted into smaller volumes. The cultures were then supplemented with either vehicle
(water) or M+6 labelled NMN (0.1 mM) and OD600 measured at time zero (before NMN), time zero (after NMN), and 140, 160 and 180 minutes after supplementation with NMN. The supernatant of cells was separated from the cells via centrifugation (5000 g for 10 minutes at
4℃) and stored immediately at -30℃. Meanwhile, the cell pellet was resuspended in cold (4°C) saline solution (0.9% NaCl) and centrifuged as above, to rinse away residual media before storage at -30℃. The OD600 was measured for each sample and used to normalise metabolite levels after LC-MS/MS analysis.
Primary hepatocyte culture
Primary hepatocytes were obtained as described previously 69. Male Sprague Dawley rats (250 grams, Animal Resources Centre, Perth, WA, Australia) were maintained on a 12:12 h day- night cycle, with water and food supplied ad libitum. Under deep non-recoverable general
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anaesthesia (75 mg/kg ketamine, 10 mg/kg xylazine, intraperitoneal administration) rats
underwent laparotomy. The portal vein was cannulated in situ and the liver perfused initially
with carbogen-saturated perfusion media (final: NaCl 138 mM, HEPES 25 mM, D-glucose 5.6
mM, KCl 5.4 mM, Na2HPO4, 0.34 mM, KH2PO4 0.44 mM, NaHCO3 4.17 mM, EDTA 0.5 mM, pH 7.4, 37°C, 25 ml/min flow rate). The inferior vena cava was cut to allow efflux. After 4 mins, the carbogen-saturated perfusion media was changed to the collagenase containing buffer
(final: NaCl 138 mM, HEPES 25 mM, D-glucose 5.6 mM, KCl 5.4 mM, Na2HPO4, 0.34 mM,
KH2PO4 0.44 mM, NaHCO3 4.17 mM, CaCl2 2 mM, collagenase II (Sigma, 15950-017), pH
7.4, 37°C, 25 ml/min flow rate) for 6 mins. The inferior vena cava was clamped at least 10
times during the collagenase digestion (preventing efflux) resulting in liver swelling that allows
a better digestion.
Following the collagenase digestion, the liver was removed and place on ice in 20 ml Williams’
Medium E (Life-technologies, Waltham, MA, USA). The hepatocytes were gently dispersed
in the medium and the cells filtered through a 100 µm cell strainer. Hepatocytes were washed
and diluted Williams’ Medium E and plated (6-well plates) at 106/2ml/well. After 4 hrs of
incubation the culture medium was changed to MOPS buffer (final, NaCl 128 mM, MOPS 23.9
mM, KCl 6 mM, MgSO4.7H2O 1.18 mM, CalCl2 1.29 mM, glucose 5 mM, BSA (FFA) 0.2 %,
pH 7.4) and cells incubated overnight. Following overnight incubation, cells were incubated in
M199 media without glutamine (Sigma M2154), supplemented with either unlabelled (Sigma)
or 15N-amide labelled glutamine (Cambridge Isotope Laboratories NLM-557) at 4 mM, in the presence or absence of NMN1 or NMN2 isotopes at 200 µM for 24 hr, following which samples were preserved for metabolomic analysis (Fig. 3).
267 APPENDIX
Preparation of NAD+ metabolite standards
NAD+ metabolites were serially diluted starting from a concentration of 100 µM to 0.39 µM.
The same volume (500 µL) of extraction buffer (acetonitrile:methanol:water) was added and
vortexed before centrifuging and transferring to new tube ready to be dried down as above. The subsequent steps were the same as preparing the tissue samples as above. All standards and samples were processed on the same day to reduce any experimental bias or variability.
Standard curves used to calculate absolute concentrations are shown on the following page
(Methods Fig. 2) and are available in supplementary raw data files.
268 APPENDIX
Above: NAD+ metabolite standard curves. (a) NMN (b) Nam (c) NAD+ (d) NR (e) Na (f) NaMN
(g) NaAD (h) NaR. Standard curves were serially diluted from 50 µM to 0.39 µM. Shown here
from 50 µM to 0.39 µM for all metabolites except for NR which is shown from 50 mM to 0.39
µM, due to the unexpectedly high concentrations of NR in the GIT (Fig. 1, 5, 6) and NaR from
100 µM to 0.39 µM.
Mass spectrometry
The LC-MS method was performed using 1260 Infinity LC System (Agilent) coupled to
QTRAP 5500 (AB Sciex) mass spectrometer. LC separation by gradient elution was
accomplished on an XBridge BEH amide column (100 mm x 2.1 mm, 3.5 µm particle size,
Waters Corporation) at room
temperature. For the mobile phase, Solvent A is 95%:5% H2O:acetonitrile containing 20 mM
ammonium acetate and 20 mM acetic acid, and solvent B is acetonitrile. The flow rate was 200
µL/min, with the percentage of solvent B set at 85% (0 min), 85% (0.1 min), 70% (10 min),
15% (13 min), 15% (17 min), 85% (17.5 min), 85% (30 min) (Supplementary Table 1).
Injection volume was 2.5 µL. Ion source was set at 350 °C and 4500 V with polarity switching.
Mass isotopologues of metabolites were acquired by MS2, using the unscheduled multiple reaction monitoring (MRM) mode with a dwell time of 40 ms. The MS parameters
(declustering potential, collision energy and cell exit potential) (Supplementary Table 2a) and
MRM transitions were calibrated based on the monoisotopic mass of chemical standards
(Supplementary Figure 11). Data processing was performed using MSConvert (version
3.0.18165-fd93202f5) and in-house MATLAB scripts. Deconvolution scripts were developed to resolve overlapping NAAD-NAD, NAR-NR and NAMN-NMN peaks using MATLAB’s
Optimisation Toolbox. Representative chromatograms are shown below.
269 APPENDIX
a Left: chromatograms represent individual peaks
(ion count) from 100µM
standard solutions at each
respective retention times for
b (a) NAM, NA, MeNAM,
NAD+ and NAAD, (b) NAMN
and NMN, (c) NAR and NR.
c
Statistical analysis for mass spectrometry
All data are presented as mean ± standard deviation (s.d.). Statistical significance was performed using a two-way ANOVA with a Sidak’s multiple comparisons test to determine differences between groups after removing outliers using the ROUT method (Q=1%). Data in
Fig. 7 were analysed by Mann-Whitney U-test between NMN isotope treated groups due to the absence of detection of labelled metabolites in animals that did not receive NMN isotopes. All statistics were performed on GraphPad Prism software (version 8.2.1). P values less than 0.05 were considered statistically significant. All data analyses are available as an .xml file available on our Mendeley data site. For in vivo experiments (Fig. 1, 5-8, Supp. Fig. 4-7), each data point
270 APPENDIX represents tissues from a separate animal, while each data point for in vitro experiments (Fig.
2, 3) represents an independent biological replicate.
DNA extraction from faecal pellets
Solid faecal pellets taken from the colonic and rectal region of the gastrointestinal tract were stored in -80℃ until further processing. DNA was extracted from frozen faecal pellets using the QIAamp® PowerFecal® DNA kit (Qiagen, Cat. No. 12830-50) according to the manufacturer’s protocol. DNA concentration was determined using the NanoDrop™
(DeNovix®, DS-11 FX) and the purity of double-stranded DNA (dsDNA) was also determined by measuring the 260/280 ratio. All DNA extracts were stored at -80℃ until further processing by 16S rRNA sequencing.
16S Sequencing
Full length 16S rRNA genes were amplified by PCR using the Oxford Nanopore 16S
Barcoding Kit (SQK-RAB204; Oxford Nanopore Technologies, Oxford, UK). Briefly, 10 ng genomic DNA, 1 µL 16S Barcode (10 µM) and 25 µL LongAmp Taq 2X Master Mix (New
England Biolabs, Ipswich, MA, USA) were combined in a 50 µL reaction for PCR on a Bio-
Rad T100TM Thermal Cycler (Bio-Rad Laboratories Pty Ltd, Hercules, CA, USA). PCR cycling condition were as follows; initial denaturation at 95 ℃ for 1 minute, 25 cycles of denaturation at 95 ℃ for 20 seconds, annealing at 55 ℃ for 30 seconds and extension at 65 ℃ for 2 minutes before a final extension at 65 ℃ for 5 minutes.
PCR products were purified as per Oxford Nanopore Technologies (ONT) protocol using
AMPure XP magnetic beads (Beckman Coulter, Indianapolis, IN) and DNA quantified using the NanoDrop™ (DeNovix®, DS-11 FX). Barcodes were pooled to a total of 100 fmol in 10
μL of 10 mM Tris-HCl, pH 8.0 with 50 mM NaCl for library loading. Sequencing was
271 APPENDIX
performed using R9.4.1 ONT Flow Cells on the MinIONTM sequencing platform and data
acquired using MinKNOW software version 19.10.1 (Oxford Nanopore Technologies).
Data Analysis for 16S sequencing
Full length 16S sequencing reads acquired from MinION runs (i.e. FAST5 data) were base-
called to fastq files using Guppy software version 3.4.4 (Oxford Nanopore Technologies). Fastq
files were demultiplexed using Porechop (https://github.com/rrwick/Porechop) and trimmed to
1400bp with Trimmomatic version 0.39 70. Reads were imported to QIIME2 for dereplication
and chimeric reads screened and filtered from the dataset. Operational taxonomic unit
clustering was completed within QIIME2 version 2019.7.0 71 at 85% similarity to account for
typical sequencing errors obtained from long-read sequencing. Taxonomy was assigned to
reads using a pre-trained classifier on the SILVA 132 16S rRNA representative sequences.
Data was imported into R version 3.6.1 with qiime2R version 0.99.13
(https://github.com/jbisanz/qiime2R) for visualisation and alpha diversity analysis using raw
and rarefied data with the phyloseq version 1.30.0 72 package. Scripts for command line processing and analysis in R available in Supplementary Materials. Sequencing data has been deposited in the NCBI database Sequence Read Archive (SRA) under accession numbers
PRJNA635359.
Figures
Labelling schemes shown in Fig. 1-4, 7 were created using BioRender.com
272 APPENDIX
SUPPLEMENTARY TABLES
Supplementary Table 1.
Time (mins) Flow rate (µl/min) Buffer A (%) Buffer B (%) 0 200 15 85 0.1 200 15 85 10 200 30 70 13 200 85 30 17 200 85 30 17.5 200 15 85 30 200 15 85 Supplementary Table 2. Liquid chromatography (LC) separation gradient
Buffer A: 95:5 (v/v) HPLC H2O:Acetonitrile (CH3CN) with 20 mM ammonium acetate
(NH4OAc) + 20mM acetic acid (CH3COOH), pH 5. Buffer B: 100% Acetonitrile (CH3CN).
273 APPENDIX
Supplementary Table 2.
Metabolite_Q1_Q3 Q1 Q3 DP CE CXP RT (m/z) (m/z) (V) (V) (V) (mins) glutamine_0_0 147 44 51 73 10 13 glutamine_1_0 148 44 51 73 10 13 glutamine_1_1 148 45 51 73 10 13 glutamine_2_1 149 45 51 73 10 13 NA_0_0 124 78 70 25 10 4 NA_1_1 125 79 70 25 10 4 NaAD_0_0 665 428 139 35 38 17 NaAD_5_0 670 428 139 35 38 17 NaAD_6_0 671 428 139 35 38 17 NaAD_1_0 666 428 139 35 38 17 NaAD_5_5 670 433 139 35 38 17 NaAD_10_5 675 433 139 35 38 17 NaAD_11_5 676 433 139 35 38 17 NaAD_6_5 671 433 139 35 38 17 NaAD_6_6 671 434 139 35 38 17 NaAD_11_6 676 434 139 35 38 17 NaAD_12_6 677 434 139 35 38 17 NaAD_7_6 672 434 139 35 38 17 NaAD_1_1 666 429 139 35 38 17 NaAD_6_1 671 429 139 35 38 17 NaAD_7_1 672 429 139 35 38 17 NaAD_2_1 667 429 139 35 38 17 NaAD_7_7 672 435 139 35 38 17 NAD_0_0 664 428 33 36 31 17 NAD_5_0 669 428 33 36 31 17 NAD_6_0 670 428 33 36 31 17 NAD_7_0 671 428 33 36 31 17 NAD_1_0 665 428 33 36 31 17 NAD_2_0 666 428 33 36 31 17 NAD_5_5 669 433 33 36 31 17 NAD_6_6 670 434 33 36 31 17 NAD_7_7 671 435 33 36 31 17 NAD_1_1 665 429 33 36 31 17 NAD_2_2 666 430 33 36 31 17 NAD_10_5 674 433 33 36 31 17 NAD_11_5 675 433 33 36 31 17 NAD_12_5 676 433 33 36 31 17 NAD_6_1 670 429 33 36 31 17 NAD_7_1 671 429 33 36 31 17 NAD_8_1 672 429 33 36 31 17
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NAD_11_6 675 434 33 36 31 17 NAD_12_6 676 434 33 36 31 17 NAD_13_6 677 434 33 36 31 17 NAM_0_0 123 80 80 30 25 2 NAM_1_1 124 81 80 30 25 2 NAM_2_1 125 81 80 30 25 2 NAM_1_0 124 80 80 30 25 2 NaMN_0_0 336 124 66 28 11 19 NaMN_5_0 341 124 66 28 11 19 NaMN_6_1 342 125 66 28 11 19 NaMN_1_1 337 125 66 28 11 19 NaR_0_0 256 124 41 27 6 10 NaR_5_0 261 124 41 27 6 10 NaR_6_1 262 125 41 27 6 10 NaR_1_1 257 125 41 27 6 10 NMN_0_0 335 123 48 24 11 19 NMN_5_0 340 123 48 24 11 19 NMN_6_1 341 124 48 24 11 19 NMN_7_2 342 125 48 24 11 19 NMN_1_1 336 124 48 24 11 19 NMN_2_2 337 125 48 24 11 19 NR_0_0 255 123 64 30 13 11 NR_5_0 260 123 64 30 13 11 NR_6_1 261 124 64 30 13 11 NR_7_2 262 125 64 30 13 11 NR_1_1 256 124 64 30 13 11 NR_2_2 257 125 64 30 13 11 Thymine-d4 129 42 -115 -52 -11 2 CSA 231 80 -170 -40 -13 2 MES 196 100 140 31 25 4
Supplementary Table 2. MRM transitions and MS parameters of NAD+ metabolites and
MRM internal standards (Thymidine d4, CSA, MES). Q1: parent ion, Q3: fragment ion, MRM: multiple reaction monitoring, DP: declustering potential, CE: collision energy, CXP: collision cell exit potential.
275 APPENDIX
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