Reduced nicotinamide mononucleotide is a new and potent NAD+precursor in mammalian cells and mice

The FASEB JournalVolume 35, Issue 4 e21456 RESEARCH ARTICLEOpen Access Reduced nicotinamide mononucleotide is a new and potent NAD+ precursor in mammalian cells and mice Rubén Zapata-Pérez, Corresponding Author Rubén Zapata-Pérez [email protected] orcid.org/0000-0003-4432-9652 Laboratory Genetic Metabolic Diseases, Amsterdam Gastroenterology, Endocrinology, and Metabolism, Amsterdam Cardiovascular Sciences, Amsterdam UMC, University of Amsterdam, Amsterdam, The Netherlands Correspondence Rubén Zapata-Pérez and Riekelt H. Houtkooper, Laboratory Genetic Metabolic Diseases, Amsterdam Gastroenterology, Endocrinology, and Metabolism, Amsterdam Cardiovascular Sciences, Amsterdam UMC, University of Amsterdam, Meibergdreef 9, Amsterdam, The Netherlands.Search for more papers by this authorAlessandra Tammaro, Alessandra Tammaro orcid.org/0000-0003-3128-5259 Pathology Department, Amsterdam UMC, University of Amsterdam, Amsterdam, The NetherlandsSearch for more papers by this authorBauke V. Schomakers, Bauke V. Schomakers Laboratory Genetic Metabolic Diseases, Amsterdam Gastroenterology, Endocrinology, and Metabolism, Amsterdam Cardiovascular Sciences, Amsterdam UMC, University of Amsterdam, Amsterdam, The Netherlands Core Facility Metabolomics, Amsterdam UMC, University of Amsterdam, Amsterdam, the NetherlandsSearch for more papers by this authorAngelique M. L. Scantlebery, Angelique M. L. Scantlebery Laboratory Genetic Metabolic Diseases, Amsterdam Gastroenterology, Endocrinology, and Metabolism, Amsterdam Cardiovascular Sciences, Amsterdam UMC, University of Amsterdam, Amsterdam, The NetherlandsSearch for more papers by this authorSimone Denis, Simone Denis Laboratory Genetic Metabolic Diseases, Amsterdam Gastroenterology, Endocrinology, and Metabolism, Amsterdam Cardiovascular Sciences, Amsterdam UMC, University of Amsterdam, Amsterdam, The NetherlandsSearch for more papers by this authorHyung L. Elfrink, Hyung L. Elfrink Laboratory Genetic Metabolic Diseases, Amsterdam Gastroenterology, Endocrinology, and Metabolism, Amsterdam Cardiovascular Sciences, Amsterdam UMC, University of Amsterdam, Amsterdam, The Netherlands Core Facility Metabolomics, Amsterdam UMC, University of Amsterdam, Amsterdam, the NetherlandsSearch for more papers by this authorJudith Giroud-Gerbetant, Judith Giroud-Gerbetant Nestlé Institute of Health Sciences, Nestlé Research, Lausanne, SwitzerlandSearch for more papers by this authorCarles Cantó, Carles Cantó Nestlé Institute of Health Sciences, Nestlé Research, Lausanne, SwitzerlandSearch for more papers by this authorCarmen López-Leonardo, Carmen López-Leonardo Department of Organic Chemistry, University of Murcia, Murcia, SpainSearch for more papers by this authorRebecca L. McIntyre, Rebecca L. McIntyre Laboratory Genetic Metabolic Diseases, Amsterdam Gastroenterology, Endocrinology, and Metabolism, Amsterdam Cardiovascular Sciences, Amsterdam UMC, University of Amsterdam, Amsterdam, The NetherlandsSearch for more papers by this authorMichel van Weeghel, Michel van Weeghel Laboratory Genetic Metabolic Diseases, Amsterdam Gastroenterology, Endocrinology, and Metabolism, Amsterdam Cardiovascular Sciences, Amsterdam UMC, University of Amsterdam, Amsterdam, The Netherlands Core Facility Metabolomics, Amsterdam UMC, University of Amsterdam, Amsterdam, the NetherlandsSearch for more papers by this authorÁlvaro Sánchez-Ferrer, Álvaro Sánchez-Ferrer Department of Biochemistry and Molecular Biology-A, University of Murcia, Murcia, SpainSearch for more papers by this authorRiekelt H. Houtkooper, Corresponding Author Riekelt H. Houtkooper [email protected] orcid.org/0000-0001-9961-0842 Laboratory Genetic Metabolic Diseases, Amsterdam Gastroenterology, Endocrinology, and Metabolism, Amsterdam Cardiovascular Sciences, Amsterdam UMC, University of Amsterdam, Amsterdam, The Netherlands Correspondence Rubén Zapata-Pérez and Riekelt H. Houtkooper, Laboratory Genetic Metabolic Diseases, Amsterdam Gastroenterology, Endocrinology, and Metabolism, Amsterdam Cardiovascular Sciences, Amsterdam UMC, University of Amsterdam, Meibergdreef 9, Amsterdam, The Netherlands.Search for more papers by this author Rubén Zapata-Pérez, Corresponding Author Rubén Zapata-Pérez [email protected] orcid.org/0000-0003-4432-9652 Laboratory Genetic Metabolic Diseases, Amsterdam Gastroenterology, Endocrinology, and Metabolism, Amsterdam Cardiovascular Sciences, Amsterdam UMC, University of Amsterdam, Amsterdam, The Netherlands Correspondence Rubén Zapata-Pérez and Riekelt H. Houtkooper, Laboratory Genetic Metabolic Diseases, Amsterdam Gastroenterology, Endocrinology, and Metabolism, Amsterdam Cardiovascular Sciences, Amsterdam UMC, University of Amsterdam, Meibergdreef 9, Amsterdam, The Netherlands.Search for more papers by this authorAlessandra Tammaro, Alessandra Tammaro orcid.org/0000-0003-3128-5259 Pathology Department, Amsterdam UMC, University of Amsterdam, Amsterdam, The NetherlandsSearch for more papers by this authorBauke V. Schomakers, Bauke V. Schomakers Laboratory Genetic Metabolic Diseases, Amsterdam Gastroenterology, Endocrinology, and Metabolism, Amsterdam Cardiovascular Sciences, Amsterdam UMC, University of Amsterdam, Amsterdam, The Netherlands Core Facility Metabolomics, Amsterdam UMC, University of Amsterdam, Amsterdam, the NetherlandsSearch for more papers by this authorAngelique M. L. Scantlebery, Angelique M. L. Scantlebery Laboratory Genetic Metabolic Diseases, Amsterdam Gastroenterology, Endocrinology, and Metabolism, Amsterdam Cardiovascular Sciences, Amsterdam UMC, University of Amsterdam, Amsterdam, The NetherlandsSearch for more papers by this authorSimone Denis, Simone Denis Laboratory Genetic Metabolic Diseases, Amsterdam Gastroenterology, Endocrinology, and Metabolism, Amsterdam Cardiovascular Sciences, Amsterdam UMC, University of Amsterdam, Amsterdam, The NetherlandsSearch for more papers by this authorHyung L. Elfrink, Hyung L. Elfrink Laboratory Genetic Metabolic Diseases, Amsterdam Gastroenterology, Endocrinology, and Metabolism, Amsterdam Cardiovascular Sciences, Amsterdam UMC, University of Amsterdam, Amsterdam, The Netherlands Core Facility Metabolomics, Amsterdam UMC, University of Amsterdam, Amsterdam, the NetherlandsSearch for more papers by this authorJudith Giroud-Gerbetant, Judith Giroud-Gerbetant Nestlé Institute of Health Sciences, Nestlé Research, Lausanne, SwitzerlandSearch for more papers by this authorCarles Cantó, Carles Cantó Nestlé Institute of Health Sciences, Nestlé Research, Lausanne, SwitzerlandSearch for more papers by this authorCarmen López-Leonardo, Carmen López-Leonardo Department of Organic Chemistry, University of Murcia, Murcia, SpainSearch for more papers by this authorRebecca L. McIntyre, Rebecca L. McIntyre Laboratory Genetic Metabolic Diseases, Amsterdam Gastroenterology, Endocrinology, and Metabolism, Amsterdam Cardiovascular Sciences, Amsterdam UMC, University of Amsterdam, Amsterdam, The NetherlandsSearch for more papers by this authorMichel van Weeghel, Michel van Weeghel Laboratory Genetic Metabolic Diseases, Amsterdam Gastroenterology, Endocrinology, and Metabolism, Amsterdam Cardiovascular Sciences, Amsterdam UMC, University of Amsterdam, Amsterdam, The Netherlands Core Facility Metabolomics, Amsterdam UMC, University of Amsterdam, Amsterdam, the NetherlandsSearch for more papers by this authorÁlvaro Sánchez-Ferrer, Álvaro Sánchez-Ferrer Department of Biochemistry and Molecular Biology-A, University of Murcia, Murcia, SpainSearch for more papers by this authorRiekelt H. Houtkooper, Corresponding Author Riekelt H. Houtkooper [email protected] orcid.org/0000-0001-9961-0842 Laboratory Genetic Metabolic Diseases, Amsterdam Gastroenterology, Endocrinology, and Metabolism, Amsterdam Cardiovascular Sciences, Amsterdam UMC, University of Amsterdam, Amsterdam, The Netherlands Correspondence Rubén Zapata-Pérez and Riekelt H. Houtkooper, Laboratory Genetic Metabolic Diseases, Amsterdam Gastroenterology, Endocrinology, and Metabolism, Amsterdam Cardiovascular Sciences, Amsterdam UMC, University of Amsterdam, Meibergdreef 9, Amsterdam, The Netherlands.Search for more papers by this author First published: 16 March 2021 https://doi.org/10.1096/fj.202001826RCitations: 29AboutSectionsPDF ToolsRequest permissionExport citationAdd to favoritesTrack citation ShareShare Give accessShare full text accessShare full-text accessPlease review our Terms and Conditions of Use and check box below to share full-text version of article.I have read and accept the Wiley Online Library Terms and Conditions of UseShareable LinkUse the link below to share a full-text version of this article with your friends and colleagues. Learn more.Copy URL Abstract Nicotinamide adenine dinucleotide (NAD+) homeostasis is constantly compromised due to degradation by NAD+-dependent enzymes. NAD+ replenishment by supplementation with the NAD+ precursors nicotinamide mononucleotide (NMN) and nicotinamide riboside (NR) can alleviate this imbalance. However, NMN and NR are limited by their mild effect on the cellular NAD+ pool and the need of high doses. Here, we report a synthesis method of a reduced form of NMN (NMNH), and identify this molecule as a new NAD+ precursor for the first time. We show that NMNH increases NAD+ levels to a much higher extent and faster than NMN or NR, and that it is metabolized through a different, NRK and NAMPT-independent, pathway. We also demonstrate that NMNH reduces damage and accelerates repair in renal tubular epithelial cells upon hypoxia/reoxygenation injury. Finally, we find that NMNH administration in mice causes a rapid and sustained NAD+ surge in whole blood, which is accompanied by increased NAD+ levels in liver, kidney, muscle, brain, brown adipose tissue, and heart, but not in white adipose tissue. Together, our data highlight NMNH as a new NAD+ precursor with therapeutic potential for acute kidney injury, confirm the existence of a novel pathway for the recycling of reduced NAD+ precursors and establish NMNH as a member of the new family of reduced NAD+ precursors. Abbreviations AK adenosine kinase AKI acute kidney injury ENT equilibrative nucleoside transporter ETC electron transport chain IR ischemia reperfusion NA nicotinic acid NAD+ nicotinamide adenine dinucleotide NADH reduced nicotinamide adenine dinucleotide NADS NAD+ synthase NAM nicotinamide NAMPT nicotinamide phosphoribosyltransferase NAPRT nicotinic acid phosphoribosyltransferase NMN nicotinamide mononucleotide NMNAT nicotinamide mononucleotide adenylyl transferase NMNH reduced nicotinamide mononucleotide NR nicotinamide riboside NRH reduced nicotinamide riboside NRK nicotinamide riboside kinase PARP poly (ADP-ribose) polymerase PPP pentose phosphate pathway PRPP phosphoribosyl pyrophosphate TECs tubular epithelial cells 1 INTRODUCTION Nicotinamide adenine dinucleotide (NAD+) and its reduced form (NADH) are ubiquitous molecules in the body, which play crucial roles in energy metabolism, as they act as hydride-accepting and hydride-donating coenzymes during mitochondrial oxidative phosphorylation.1 Apart from its role as a redox cofactor, during the last decade NAD+ has arisen as the critical substrate for a number of protein families, such as the sirtuin (SIRT) family of protein deacetylases,2 poly(ADP-ribose)polymerases,3 and ADP-ribose cyclases.4 Through their downstream actions, these proteins participate in more than 500 enzymatic reactions and regulate almost all major biological processes in cells.5 This continuous enzymatic utilization of NAD+ is counterbalanced via de novo synthesis from dietary tryptophan, or through its salvage from precursors. In the Preiss-Handler pathway, the NAD+ precursor nicotinic acid (NA) is converted into NAD+ in a three-step enzymatic process led by the nicotinic acid phosphoribosyltransferase (NAPRT), the nicotinamide mononucleotide adenylyl transferases (NMNATs), and the NAD+ synthase (NADS). Another recycling pathway comprises intracellular nicotinamide (NAM) phosphoribosylation or nicotinamide riboside (NR) phosphorylation into nicotinamide mononucleotide (NMN), a process carried out by nicotinamide phosphoribosyltransferase (NAMPT) or nicotinamide riboside kinases (NRKs), respectively. NMN is then directly converted to NAD+ by the NMNATs.6 When given externally, NMN can also act as an NAD+ precursor. To achieve this, NMN first needs to be converted extracellularly to NR by the ectoenzyme 5′-nucleotidase CD73, after which NR is transported into the cell by the equilibrative nucleoside transporters (ENTs) and metabolized to NAD+ via NRKs.7, 8 Very recently, it has also been reported that NMN can be incorporated into cells via the Slc12a8-specific transporter, at least in mouse small intestine.9 The role of NAD+ in the activity of enzymes controlling major metabolic processes, together with reports supporting that decreased cellular NAD+ contributes to metabolic disturbances,10 have renewed the interest in strategies to increase NAD+ bioavailability to combat disease. In fact, NAD+ repletion and the subsequent activation of sirtuins leads to key biochemical and clinical improvements, such as enhanced mitochondrial biogenesis,11-14 protection against fatty acid-induced liver disease15 and diabetes,11, 12 or reduced neurodegeneration16 in a variety of animal models. NAD+ homeostasis also plays a major role in kidney health and in the ability of the renal tubule to resist stressors.17 In fact, during ischemic renal injury, NAD+ consumption by poly (ADP-ribose) polymerases (PARPs) is accelerated,18 leading to NAD+ decline in renal tissue. NAD+ replenishment through administration of the NAD+ precursor NMN has been proven effective in ameliorating tubular damage induced by ischemia-reperfusion (IR) injury and the nephrotoxic drug cisplatin in aged mice.19 These results have turned attention to the use of NAD+ precursors for combatting metabolic disease in humans. NA and NAM have downsides, however. The former induces flushing triggered by NA binding to the GPR109A receptor,20 and NAM can act as a sirtuin inhibitor, which could limit its intended activation of these enzymes. Therefore, NMN and NR have arisen as attractive alternatives to NA or NAM, since they effectively raise NAD+ concentrations in mouse tissues without undesired adverse target effects.11, 12, 21, 22 For this reason, several clinical studies have been initiated with NR. Some studies23-28 with NR have shown that this compound is well tolerated up to 2 g per day during 12 weeks, while the first clinical trials with NMN are still ongoing (NCT03151239, UMIN000021309, UMIN000030609, and UMIN000025739). Yet, NMN and NR supplementation have some limitations of their own, including maximal NAD+-enhancing effects of around 2-fold, the need of high doses (from 200 to 1000 mg/kg per day) to achieve beneficial effects in animal models,11-13, 22, 29 and the rapid degradation in plasma to NAM, at least in the case of NR.30 Moreover, although an increase in NAD+ levels in whole blood has been detected upon NR administration in humans,24, 27 supplementation with this precursor has failed to increase NAD+ in other tissues, such as muscle biopsies, even after 1 g administration during 6 weeks.31 This inefficacy in raising NAD+ might explain why NR has no apparent effect on total energy expenditure, blood glucose or insulin sensitivity in humans.28 To overcome the limitations of the current repertoire of NAD+ enhancers, other molecules with a more pronounced effect on the NAD+ intracellular pool are desired. This has stimulated us to investigate the use of the reduced form of nicotinamide mononucleotide (NMNH) as an NAD+ enhancer. There is very scarce information about the role of this molecule in cells. In fact, only one enzymatic activity has been described to produce NMNH. This is the NADH diphosphatase activity of the human peroxisomal Nudix hydrolase hNUDT1232 and the murine mitochondrial Nudt13.33 It has been postulated that, in cells, NMNH would be converted to NADH via nicotinamide mononucleotide adenylyl transferases (NMNATs).34 However, both NMNH production by Nudix diphosphatases and its use by NMNATs for NADH synthesis have only been described in vitro using isolated proteins, and how NMNH participates in cellular NAD+ metabolism remains unknown. In the present study, we design and develop a new method for the purification of NMNH at scale, and explore the role of this molecule in NAD+ metabolism. We show that NMNH is effectively metabolized to NAD+ in mammalian cells, and confirm its NAD+ synthesis route is NRK and NAMPT-independent. We also investigate the therapeutic potential of NMNH, showing that it can protect renal proximal tubular epithelial cells from hypoxia/reoxygenation-induced injury, a crucial event in ischemic acute kidney injury (AKI),35 by accelerating processes involved in tubular regeneration.36 Finally, we explore the in vivo effects of NMNH administration in mice and demonstrate that this new precursor effectively raises NAD+ levels in blood and a variety of tissues, including kidney, to a greater extent than NMN when used at the same concentration. These results corroborate that reduced NAD+ precursors can act as very potent NAD+ enhancers, and open doors for a new generation of highly efficient NAD+-boosting molecules that could aid in overcoming the limitations of the current set of NAD+ enhancers. 2 MATERIALS AND METHODS Enzyme preparation and NMNH synthesis The pyrophosphatase from Escherichia coli strain K12 (EcNADD, Uniprot code: P32664) was cloned into pET24b vector and expressed in E coli Rosetta 2 DE3 pLysS at 25°C and 0.25 mM of isopropyl-β-thiogalactopyranoside (IPTG, Sigma-Aldrich, St. Louis, MO, USA) for 16 hours. Protein purification was performed in a HiTrap IMAC column (GE Healthcare) in 50 mM of Tris-HCl pH 7.5 containing 300 mM of NaCl in a gradient from 10 to 250 mM of imidazole followed by desalting in a HiPrep desalting column (GE Healthcare, Chicago, IL, USA). Conversion of NADH into NMNH and AMP was achieved by incubation of 50 µg/mL EcNADD with 5 mM NADH in 50 mM Tris HCl pH 8.0 containing 0.5 mM MnCl2 in a total volume of 50 mL for 45 minutes at 37°C. NADH conversion was checked by HPLC using a reverse-phase C18 250 × 4.6 mm column (Phenomenex, Torrance, CA, USA) and a mobile phase consisting of 20 mM ammonium acetate pH 6.9 running at 1 mL/min for 15 minutes. Under these chromatographic conditions, retention times for NMNH and AMP were 5.5 and 7 minutes, respectively. NMNH purification Chromatographic separation of NMNH from AMP was achieved by C18 reverse-phase chromatography. Fractions containing pure NMNH were evaporated until dryness in a rotary evaporator. The resulting powder was redissolved and desalted by size exclusion chromatography in pure water. After the final desalting step, samples were freeze dried to obtain NMNH as an amorphous yellow powder. Identification of NMNH by nuclear magnetic resonance The proton and 31P NMR spectra were recorded at 25°C on a Bruker (Billerica, MA, USA) Avance 400 (400 MHz). 1H chemical shifts were reported in ppm relative to the resonance of HOD (δ = 4.8 ppm) and 31P chemical shifts were externally referenced to 85% of H3PO4. J values are given in Hz. Cell culture and supplementation with NAD+ precursors AML12, HepG2, SY5Y, HeLa cells, and human skin fibroblasts were cultured in 12-well plates in Dulbecco's modified Eagle's medium (DMEM, Life Technologies, Carlsbad, CA, USA) supplemented with 10% of fetal bovine serum (FBS, BioWhittaker, Basel, CH), 100 U/mL of penicillin, and 10 mg/mL of streptomycin (Life Sciences, Waltham, MA, USA). T37i cells were cultured and differentiated as previously described.37, 38 Conditionally Immortalized Proximal Tubular Epithelial Cells (IM-PTECs) were generated as previously described39 and cultured at 33°C in HK2 medium40 supplemented with 10 ng/mL IFN-γ (ProSpec, Rehovot, IL, USA) and maintained at 37°C without IFN-γ for an additional week before the start of the assay, resulting in loss of SV40 expression.41 Otherwise indicated, supplementation with PBS (vehicle), NMN (Carbosynth, Compton, UK), or NMNH was made at the concentrations and times indicated in FBS-free medium. Chemical inhibition of the different enzymes involved in the recycling of NAD+ precursors was made using the following reagents and concentrations: (E)-N-[4-(1-benzoylpiperidin-4-yl)butyl]-3-(pyridin-3-yl)acrylamide (FK866; 2 µM; Sigma-Aldrich), adenosine-5′-[(α,β)-methylene]diphosphate (AOPCP; 500 µM; Jena Biosciences, Jena, DE), dipyridamole (DIPY; 20 µM; Sigma-Aldrich), 5-(3-Bromophenyl)-7-[6-(4-morpholinyl)-3-pyrido[2,3-d]pyrimidin-4-amine dihydrochloride (ABT702; 10 µM; Tocris Biosciences, Bristol, UK), and gallotannin (100 µM; Sigma-Aldrich). Cells were incubated for 1 hour in the presence of the corresponding inhibitor prior to supplementation with the indicated concentrations of NMNH or PBS (vehicle). Viability assay AML12 cells were cultured in DMEM in a 96-well plate and supplemented with different concentrations of NMN or NMNH. After 24 hours, cells were washed with PBS and 150 µL of MTS reagent (Abcam, Cambridge, UK) was added to each well. After incubation at 37°C for 4 hours, absorbance was measured on a spectrophotometer at 490 nm. Enzymatic cyclic assay for quantitative NAD+ determination For NAD+ extraction, cells were washed twice with PBS, quenched with 400 µL of 2 M HClO4 and transferred to 1.5 mL tubes. For tissues, 3-4 mg of freeze-dried tissue or 10 µL of blood were resuspended in 400 µL of 2 M HClO4 and disrupted using a TissueLyser (Qiagen, Venlo, NL) for 5 minutes at 30 pulses per second. After centrifugation at 16 000 g for 5 minutes, 100 µL of the acidic supernatant was neutralized by addition of 150 µL 2 M/0.6 M KOH/MOPS and centrifuged again to remove precipitated salts. NAD+ content was determined using an enzymatic spectrophotometric cycling assay based on the coupled reaction of malate and alcohol dehydrogenases, as previously described.42 Mass spectrometry measurements PBS-washed cells or 5-6 mg of freeze-dried tissue were metabolically quenched using ice-cold methanol (500 µL) and diluted with Milli-Q water (500 µL) containing internal standards, D5-glutamine, D5-phenylalanine, adenosine-15N5-monophosphate, adenosine-15N5-triphosphate, and guanine-15N5-triphosphate (5 μM each). Phase separation was performed by chloroform addition (1 mL) followed by thorough mixing. The homogenate was centrifuged at 16 000 g for 5 minutes at 4°C and the polar upper phase was transferred to a new 1.5 mL tube. Samples were then dried in a vacuum concentrator and pellets dissolved in 100 µL of methanol/water (6/4; v/v). Metabolite analysis was carried out in an Acquity UPLC system (Waters, Milford, MA, USA) coupled to an Impact II Ultra-High Resolution Qq-Time-Of-Flight mass spectrometer (Bruker). Chromatographic separation of the compounds was achieved using a SeQuant ZIC-cHILIC column (PEEK 100 × 2.1 mm, 3 µm particle size; Merck, Kenilworth, NJ, USA) at 30°C. The LC method consisted in a gradient running at 0.25 mL/min from 100% mobile phase B (9:1 acetonitrile:water with 5 mM ammonium acetate pH 8.2) to 100% mobile phase A (1:9 acetonitrile:water with 5 mM ammonium acetate pH 6.8) in 28 minutes, followed by a re-equilibration step at 100% B of 5 minutes. MS data were acquired both in negative and positive ionization modes over the range of m/z 50-1200. Data from full-scan MS mode was analyzed using Bruker TASQ software (Version 2.1.22.1 1065). All reported metabolite intensities were normalized to total protein content in samples, determined using a Pierce BCA Protein Assay Kit, as well as to internal standards with comparable retention times and response in the MS. For MS/MS mode, the parent mass in positive mode was selected at m/z 335.1 ± 0.5 for NMN and m/z 337.1 ± 0.5 for NMNH both at 10 eV collision energy using ultrapure nitrogen as collision gas. Parent and subsequent daughter fragments were analyzed using Bruker Compass DataAnalysis (Version 5.1, Build 201.2.4019). Hypoxia/reoxygenation assays In order to establish an in vitro model of tubular injury induced by hypoxia/reoxygenation, IM-PTECs were grown under oxygen-deficient conditions (1% O2 and 5% CO2 at 37°C) for 48 hours using a hypoxic incubator (HypOxystation H35; Don Whitley Scientific, Bingley, UK). Normoxic control cells were cultured under standard conditions (21% O2 and 5% CO2 at 37°C). After 48 hours, both hypoxic and normoxic cells received fresh HK2 medium, were cultured for an additional 24 hours under standard conditions (reoxygenation) and supplemented with the corresponding compounds, as indicated in the figure legends. Mitochondrial superoxide measurements To examine the effect of NAD+ boosting during hypoxia/reoxygenation on mitochondrial superoxide production, IM-PTECs were labeled with 5 µM of MitoSox Red (Thermo Fisher Scientific, Waltham, MA, USA) in HBSS supplemented with calcium and magnesium (Gibco, Waltham, MA, USA) for 10 minutes at 37°C. Relative fluorescence was measured by FACS. BrdU incorporation assay To analyze the cell cycle, TECs subjected to hypoxia/reoxygenation conditions were incubated for 1 hour with 20 mM of bromodeoxyuridine (BrdU), and subsequently fixed in ice-cold ethanol. Cells were then incubated with 0.4 mg/mL of pepsin and 0.2 mM of HCl for 30 minutes at 37°C, followed by incubation with 2 M HCl for 25 minutes at 37°C. After washing, cells were stained for 30 minutes with anti BrdU-FITC (clone B44, BD Biosciences, San Jose, CA, USA) in PBS supplemented with 0.05% of Tween-20 and 0.5% of BSA. Finally, cells were treated with 500 µg/mL RNAse-A (Bioke, Leiden, NL) and stained with 0.1 µM of TO-PRO-3-iodide (Invitrogen, Life technologies, Carlsbad, CA, USA) for 15 minutes at 37°C. Cells were acquired on a LSRFortessa cell analyzer (BD Biosciences). Cell cycle distribution was analyzed with the FlowJo 7.6 software (Tree Star, Ashland, OR, USA). The percentage of cells in S phase was determined by BrdU uptake. Western blotting TECs were lysed in RIPA buffer (50 mM Tris-HCl pH 7.5, 150 mM NaCl, 2 mM EDTA, 1% deoxycholic acid, 1% NP-40, 4 mM sodium orthovanadate, 10 mM sodium fluoride), supplemented with protease and phosphatase inhibitors. Lysates were loaded onto a 4%-12% pre-casted gel and blotted onto a PVDF membrane. The antibodies used were β-actin (Sigma-Aldrich) and cleaved caspase-3 (Cell signaling, Danvers, MA, USA). mRNA extraction and analysis Total RNA was extracted using TRIzol-reagent (Sigma-Aldrich). cDNA was synthesized using M-MLV reverse transcriptase and oligo-dT primers (Table S1). Transcript analysis was performed by real-time quantitative PCR on a Roche Light Cycler 480 using SYBR green master mix (Bioline, London, UK). Relative expression was analyzed using LinRegPCR.43 Gene expression was normalized to murine peptidylprolyl isomerase A (Ppia) as a housekeeping gene. In vivo study Animal experiments were performed according to national Dutch and EU ethical guidelines and approved by the local animal experimentation committee under license AVD1180020186906. All the experiments were performed in male C57BL/6N mice at 11 weeks old. Mice were kept in a temperature- and humidity-controlled environment under a 12-hour light/dark cycle with free access to food and water. Twenty-four mice were randomly assigned to three groups and intraperitoneally (IP) injected with saline, NMN or NMNH at 250 mg/kg. Blood was collected from the lateral saphenous vein at 1, 4, and 20 hours after the first IP injection. After 20 hours, another bolus injection was administered and 4 hours after this last injection, mice were sacrificed under isoflurane, and blood and tissues collected and snap frozen in liquid nitrogen until NAD+ determination and mass spectrometry measurements. 3 RESULTS Enzymatic synthesis and purification of NMNH Since NMNH is not commercially available, we developed a preparation method of the compound. To do so, we took advantage of the high activity of the NAD+ pyrophosphatase from Escherichia coli (EcNADD) to cleave NADH into NMNH and AMP44 (Figure 1A). Indeed, with 2.5 mg of the recombinant enzyme we were able to fully convert 175 mg of NADH into NMNH and AMP in 45 minutes at 37°C (Figure S1A). The two molecules were then successfully separated using C18 chromatography (Figure S1B), desalted and freeze dried until an amorphous yellow powder was obtained. The final yield of the process was approximately 70%. FIGURE 1Open in figure viewer NMNH synthesis and characterization. A, Synthesis and purification of NMNH from NADH. B, Linear curve of NMNH concentration vs absorbance at 340 nm. C, 1H and 31P NMR data for NMNH. 1H NMR (400 MHz, D2O): δ 2.96 (br s, 2H, H4A + H4B), 3.75 (br s, 2H, H5A' + H5B'), 3.99 (bs s, 1H, H3'), 4.13-4.16 (m, 1H, H2'), 4.22-4.26 (m, 1H, H4'), 4.77-4.79 (m, 1H, H1'), 4.86-4.94 (m, 1H, H5), 6.13 (d, 1H, J = 8.0 Hz, H6), and 7.06 (s, 1H, H2). Inset: 31P NMR data (161 MHz, D2O): δ 3.83 (D and E) MS/MS analysis of pure NMN (D) and NMNH (E) The chemical structure and mass of NMNH were confirmed by UV spectrophotometry, nuclear magnetic resonance (NMR) and mass spectrometry. NMNH was shown to be a fluorescent molecule, in contrast to NMN, with a molar extinction coefficient of 5160 M−1 cm−1 at 340 nm (Figure 1B), a value that is very similar to that found for the reduced form of nicotinamide riboside (NRH).45 Structural analysis of NMNH was performed by 1H and 31P NMR (400 MHz, D2O; Figure 1C), matching expectations and showing the two characteristic H4 hydrogens of NMNH at 3 ppm. NMNH exact mass and fragmentation pattern were analyzed by mass spectrometry in positive ionizatio