Published: August 13, 2019

Introduction {#sec1}
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Aging is characterized by a decline in metabolic and physiological functions of all organs within the body. A hallmark feature of aging is the progressive loss of skeletal muscle mass and function that can progress to sarcopenia, which is associated with significant morbidity and mortality and substantial healthcare costs ([@bib38], [@bib60]). Exercise is considered a frontline modality to combat age-related muscle decline ([@bib13]). However, nutritional strategies may also offer an effective countermeasure to age-associated morbidities and promote healthy muscle aging ([@bib6]).

Nicotinamide adenine dinucleotide (NAD^+^) homeostasis is critical to cell and organismal function. In addition to its classical role in redox metabolism, NAD^+^ is a substrate for enzymes such as sirtuins, poly-ADPribose polymerases (PARPs), and cyclic ADPribose synthetases that regulate key cellular processes of energy metabolism, DNA damage repair, and calcium signaling ([@bib71]). Improving NAD^+^ availability via the supplementation of the NAD^+^ precursor nicotinamide riboside (NR) ([@bib5], [@bib63]) has emerged as a potential strategy to augment tissue-specific NAD^+^ homeostasis and improve physiological function ([@bib22]). A range of physiological stresses associated with the depletion of NAD^+^ and/or nicotinamide adenine dinucleotide phosphate hydrogen (NADPH) have been ameliorated with NR supplementation in mice, including prevention of noise-induced hearing loss ([@bib7]), resistance to weight gain ([@bib9]), reduction of blood glucose, hepatic steatosis and neuropathy on a high-fat diet ([@bib64]), improvement of cardiac function in genetic cardiomyopathy ([@bib18]), and prevention of cortical neuronal degeneration ([@bib67]). Depletion of the enzyme nicotinamide phosphoribosyltransferase (NAMPT), rate-limiting for NAD^+^ biosynthesis, in mouse skeletal muscle severely diminishes NAD^+^ levels and induces sarcopenia. Oral repletion of NAD^+^ with NR in this model rescued pathology in skeletal muscle in a cell-autonomous manner ([@bib25]). However, recent data in mice tracing NAD^+^ fluxes questioned whether oral NR has the ability to access muscle ([@bib45]). Thus, whether oral NR can augment the human skeletal muscle NAD^+^ metabolome is currently unknown.

A decline in NAD^+^ availability and signaling appears to occur as part of the aging process in many species ([@bib27], [@bib52]), though there is a paucity of data to confirm that this is the case in human aging. NR and nicotinamide mononucleotide (NMN) are reported to extend life spans ([@bib72]) and enhance metabolism in aged mice ([@bib50]). To date, NR supplementation studies in humans have focused on cardiovascular ([@bib46]), systemic metabolic ([@bib20]), exercise ([@bib21]), and safety ([@bib12]) end-points, but have not addressed advanced aging, tissue metabolomic changes, or effects on muscle metabolism and function.

Herein, we set out to study if oral NR is available to aged human skeletal muscle and whether potential effects on muscle metabolism can be detected. We conducted a 21-day NR supplementation intervention in a cohort of 70--80-year-old men in a placebo-controlled, double-blind, crossover trial. We demonstrate that NR augments the skeletal muscle NAD^+^ metabolome, inducing a gene expression signature suggestive of downregulation of energy metabolism pathways, but without affecting muscle mitochondrial bioenergetics or metabolism. Additionally, we show that NR suppresses specific circulating inflammatory cytokine levels.

Results {#sec2}
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Oral NR Is Safe and Well-Tolerated in Aged Adults {#sec2.1}
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Twelve aged (median age of 75 years) and marginally overweight (median BMI of 26.6 kg/m^2^; range 21--30), but otherwise healthy, men were recruited and orally supplemented with 1-g NR per day for 21 days in a placebo-controlled, randomized, double-blind, crossover design, with 21 days' washout period between phases. Baseline characteristics of participants are included in [Table S1](#mmc1){ref-type="supplementary-material"}. NR chloride (Niagen) and a placebo were provided as 250-mg capsules (ChromaDex), and subjects were instructed to take two in the morning and two in the evening. All participants completed the study visits (5 in total) and assessments according to protocol ([Figure S1](#mmc1){ref-type="supplementary-material"}). Visit 1 was a screening and enrollment visit, while visit 4 was after the washout period, and only fasting blood and 24-h urine were collected. The protocol design for visits 2, 3, and 5 included muscle biopsy, fasting blood analyses, glucose tolerance test, muscle arterio-venous difference technique, venous occlusive plethysmography, and indirect calorimetry analysis ([Figure S1](#mmc1){ref-type="supplementary-material"}). NR was well tolerated, and screening for a range of hematological and clinical biochemistry safety parameters (including renal, liver, and thyroid functions) revealed no adverse effects ([Table S2](#mmc1){ref-type="supplementary-material"}). No clinical adverse events were reported during the intervention in either phase. Of note, four participants (33.3%), blinded to the intervention arm, self-reported a noticeable increase in libido while on NR. There were no such reports while on the placebo.

Oral NR Augments the Skeletal Muscle NAD^+^ Metabolome {#sec2.2}
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To assess the effects of NR supplementation on NAD^+^ metabolism, we used a targeted liquid chromatography-mass spectrometry (LC-MS/MS) method ([@bib62]) to quantify the NAD^+^ metabolome in skeletal muscle, whole venous blood, and urine. We examined the NAD^+^ metabolome in skeletal muscle biopsies from all participants in a fasted state at baseline and after the NR and placebo phases, 14 h after the last dose and prior to the physiological assessments. Samples were collected 14 h after the last dose so participants could attend in a fasted state, as well as to evaluate the effects of longer-term NR administration rather than those of an acute dose. Fourteen metabolites were measured in muscle extracts ([Figures 1](#fig1){ref-type="fig"} and [S2](#mmc1){ref-type="supplementary-material"}A; [Table S3](#mmc2){ref-type="supplementary-material"}). NR was detectable in muscle but was not elevated in the NR supplementation period (NR 1.4 pmol/mg μM versus placebo 1.25 pmol/mg; p = 0.23). Consistent with nicotinic acid adenine dinucleotide (NAAD) as a highly sensitive biomarker of NR supplementation and an enhanced rate of NAD^+^ synthesis ([@bib63]), we found that oral NR resulted in a 2-fold increase in muscle NAAD (NR 0.73 pmol/mg versus placebo 0.35 pmol/mg; p = 0.004), without an increase in NAD^+^ (NR 210 pmol/mg versus 197 pmol/mg; p = 0.22). NR supplementation did not affect muscle nicotinamide (NAM) (NR 92.0 pmol/mg versus placebo 86.5 pmol/mng; p = 0.96). However, remarkably, we detected 5-fold increases in the products of NAM methylation clearance pathways; N-methyl nicotinamide (MeNAM; NR 1.45 pmol/mg versus placebo 0.35 pmol/mg; p = 0.006), N1-methyl-2-pyridone-5-carboxamide (Me-2-py; NR 6.6 pmol/mg versus placebo 1.1 pmol/mg; p \< 0.001), and N1-methyl-4-pyridone-5-carboxamide (Me-4-py; NR 1.6 pmol/mg versus placebo 0.3 pmol/mg; p \< 0.001) ([Figures 1](#fig1){ref-type="fig"} and [S2](#mmc1){ref-type="supplementary-material"}A; [Table S3](#mmc2){ref-type="supplementary-material"}).Figure 1NR Augments the Human Skeletal Muscle NAD^+^ MetabolomeSchematic representation of nicotinamide riboside (NR) metabolism within the nicotinamide adenine dinucleotide (NAD^+^) metabolome, accompanied by observed levels of metabolites measured using LC-MS/MS in skeletal muscle, whole blood, and urine at baseline and after each of the NR and placebo periods. NAD^+^ metabolomics data at the end of the washout period are shown in [Table S3](#mmc2){ref-type="supplementary-material"}. Skeletal muscle data were normalized to the weight of the muscle pellet used for extraction. Urine data were normalized to urinary creatinine. Other metabolites are shown in [Figure S2](#mmc1){ref-type="supplementary-material"}. Data are obtained from 12 participants at each phase and presented as mean ± SEM. Significance was set at p \< 0.05 using paired t tests and represents the differences between NR and the placebo and between NR and baseline. The absence of significance symbols indicates a lack of statistical significance. BLQ, below limit of quantification; NMN, nicotinamide mononucleotide; NAAD, nicotinic acid adenine dinucleotide; NAM, nicotinamide; NAMOx, nicotinamide N-oxide; MeNAM, N-methyl nicotinamide; Me-2-py, N1-Methyl-2-pyridone-5- carboxamide.

In the blood, we measured 15 metabolites from each participant at baseline and following each of the NR, placebo, and washout periods ([Figures 1](#fig1){ref-type="fig"} and [S2](#mmc1){ref-type="supplementary-material"}B; [Table S3](#mmc2){ref-type="supplementary-material"}). NR was also detectable in the blood but was not increased, compared to the placebo at 14 h after the last dose of NR (NR 0.16 μM versus placebo 0.15 μM; p = 0.31). This is expected, as the predicated Cmax for NR is approximately 3 h ([@bib1]). NR increased the concentrations of NAD^+^ \>2-fold (NR 47.75 μM versus placebo 20.90 μM; p \< 0.001) and NMN 1.4-fold (NR 1.63 μM versus placebo 1.13 μM; p \< 0.001). A recent study reported that oral NR is rapidly metabolized in the liver to NAM, which can enhance tissue NAD^+^ metabolomes ([@bib45]). However, chronic NR supplementation did not elevate NAM in the blood (NR 10.60 μM versus placebo 9.50 μM; p = 0.41). Again, NAM urinary clearance pathways were highly active following NR, with marked excess of MeNAM (NR 0.66 μM versus placebo 0.10 μM; p \< 0.001), Me-2-py (NR 7.69 μM versus placebo 1.44 μM; p \< 0.001), and Me-4-py (NR 3.82 μM versus placebo 0.48 μM; p \< 0.001) ([Figures 1](#fig1){ref-type="fig"} and [S2](#mmc1){ref-type="supplementary-material"}B; [Table S3](#mmc2){ref-type="supplementary-material"}). NR elevated blood NAAD levels by 4.5-fold (NR 0.18 μM versus placebo 0.04 μM; p \< 0.001).

Urinary NAD^+^ metabolomics showed that NR was detectable and increased with NR supplementation (NR 41.5 μmol/mol creatinine versus placebo 31.7 μmol/mol creatinine; p = 0.02) ([Figure 1](#fig1){ref-type="fig"}). Furthermore, a near-20-fold increase in nicotinic acid riboside (NAR; NR-185.5 μmol/mol creatinine versus placebo-10.3 μmol/mol creatinine; p = 0.001) was observed. This observation may support the suggestion that NR supplementation leads to retrograde production of NAAD, nicotinic acid mononucleotide (NAMN), and NAR ([@bib63]). However, direct NR transformation into NAR cannot be excluded. Unlike muscle and blood, NAM was elevated in the urine 2.5-fold (NR-282 μmol/mol creatinine versus placebo-106.5 μmol/mol creatinine; p = 0.004). These data establish the extent and breadth of changes to NAD^+^ metabolites in human muscle, blood, and urine after NR supplementation. The data indicate that oral NR greatly boosts the blood NAD^+^ metabolome without an increase in NAM, increases muscle NAD^+^ metabolism, and leads to the disposal of urinary clearance products.

Oral NR Results in Downregulation of Gene Sets Associated with Energy Metabolism in Skeletal Muscle {#sec2.3}
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We next assessed NR-mediated transcriptional changes in skeletal muscle. RNA sequencing followed by differential gene expression (DGE) analysis of muscle biopsies from the 12 participants revealed 690 upregulated and 398 downregulated genes between baseline and NR supplementation at p value \< 0.05 ([Figure 2](#fig2){ref-type="fig"}A; [Table S4](#mmc3){ref-type="supplementary-material"}). Using gene annotation analysis (gene set enrichment analysis \[GSEA\]) ([@bib51], [@bib61]), we examined the enrichment of genes that belong to known molecular pathways in our list of up- or downregulated genes. Our results suggest that genes significantly downregulated with NR supplementation were enriched in pathways relating to energy metabolism, including those of glycolysis, tricarboxylic acid (TCA) cycle, and mitochondria ([Figure 2](#fig2){ref-type="fig"}B; [Table S5](#mmc4){ref-type="supplementary-material"}). This is consistent with the recent discovery that oral NR depresses mitochondrial membrane potential while improving blood stem cell production in mice ([@bib66]).Figure 2NR Supplementation Induces a Transcriptional Signature in Human Skeletal Muscle(A) Differential gene expression analysis on baseline and NR-treated muscle samples (n = 12 at each phase). Volcano plot of differential gene expression between baseline and NR treated human muscle samples. Fold change (Log2, x axis) of gene expression is plotted against p value for differential gene expression (--Log10, y axis). Colored dots represent Ensembl genes that are either upregulated (in orange) or downregulated (in blue) upon NR supplementation at a p value \< 0.05.(B and C) Gene Ontology analysis of significantly dysregulated genes upon NR supplementation for (B) downregulated genes and (C) upregulated genes. Gene Ontology analysis was performed using GSEA. Bars represent the p value (--Log10) of overlap from hypergeometric distribution.(D) Gene set enrichment analysis (GSEA) suggests that genes belonging to the gene set "glycolysis" are downregulated upon NR supplementation. The normalized enrichment score (NES) and nominal p value are presented on the top-left corner of the graph.(E) As in (D), but for genes involved in the TCA cycle.(F) As in (D), but for genes involved in the gene set "mitochondria."(G) A qPCR analysis of a select panel of downregulated genes identified through differential gene expression analysis. *GAPDH* was used as housekeeping gene. Error bars represent SEM (n = 12).(H) As in (G), but for NAD^+^ pathway-related genes.(I) Quantification of phosphoglycerate kinase 1 (PGK1), phosphoglucomutase 1 (PGM1), and pyruvate kinase M1 (PKM1) proteins using immunoblotting assay. Tubulin was used as a loading control.Data are obtained from 12 participants at each phase and wherever relevant are presented as mean ± SEM. Significance was set at p \< 0.05. The absence of significance symbols indicates a lack of statistical significance.

Pathways upregulated upon NR supplementation prominently belonged to Gene Ontology categories such as cell adhesion, actin cytoskeleton organization, and cell motility ([Figure 2](#fig2){ref-type="fig"}C). This supports a previously identified role for the NAD^+^-generating enzyme NR kinase 2b (Nrk2b) in zebrafish skeletal muscle cell adhesion ([@bib29]).

We next examined all the genes that belonged to the glycolysis, mitochondrial, and TCA cycle pathways and found that they were predominantly downregulated following NR supplementation, whereas 10 control gene sets of the same size and expression level were not ([Figures 2](#fig2){ref-type="fig"}D--2F and [S3](#mmc1){ref-type="supplementary-material"}A). Similarly, we found that the genes belonging to the Gene Ontology terms actin filament-based process, cell motility, and biological cell adhesion were mainly upregulated upon NR supplementation ([Figures S3](#mmc1){ref-type="supplementary-material"}B and S3C).

In agreement with the DGE analysis, quantitative real-time PCR showed downregulation of selected genes involved in energy metabolism ([Figure 2](#fig2){ref-type="fig"}G). We found no changes in the transcript levels of key genes involved in NAD^+^ metabolism, corroborating the DGE analysis ([Figure 2](#fig2){ref-type="fig"}H). We also verified some of the upregulated targets by qPCR ([Figure S3](#mmc1){ref-type="supplementary-material"}D) and undertook some immunoblotting validation ([Figure S3](#mmc1){ref-type="supplementary-material"}D).

As it has previously been shown that NR increases glycolysis in mouse cardiac cells ([@bib18]), and because our data do not support an NR-mediated transcriptional upregulation of glycolysis related genes, we examined protein expression levels of glycolytic enzymes in our muscle biopsies and show them to be unchanged after NR ([Figure 2](#fig2){ref-type="fig"}I).

Three Weeks of Oral NR Does Not Alter Skeletal Muscle Mitochondrial Bioenergetics or Hand-Grip Strength {#sec2.4}
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Several preclinical studies suggest that NR enhances mitochondrial energy programs in skeletal muscle ([@bib9], [@bib25]) through mechanisms that involve redox and sirtuins activation. Therefore, we undertook a detailed assessment of muscle mitochondrial respiration in biopsies after NR supplementation using high-resolution respirometry, the gold standard method for the *ex vivo* assessment of mitochondrial function. No differences were detected between the NR and placebo groups in skeletal muscle complex I- and complex II-mediated oxidative phosphorylation and maximal respiratory capacity, with ([Figure 3](#fig3){ref-type="fig"}A) and without ([Figure 3](#fig3){ref-type="fig"}B) the prior addition of the fatty acid conjugate octanoyl-carnitine. In line with this, the activity of citrate synthase, commonly used as a quantitative measure of mitochondrial content ([@bib40]) ([Figure 3](#fig3){ref-type="fig"}C), and mitochondrial copy number (mtDNA) ([@bib54]) ([Figure 3](#fig3){ref-type="fig"}D) were unchanged by NR supplementation. Similarly, levels of skeletal muscle biopsy mitochondrial resident proteins, directly involved in the electron transport chain, were unaltered upon NR supplementation ([Figure 3](#fig3){ref-type="fig"}E). We then tested whether the NR-driven increase in the NAD^+^ metabolome translates into higher sirtuin-mediated deacetylation activity, and we performed western blotting to assess pan-acetylation status, but again did not detect NR-mediated changes to muscle protein acetylation ([Figure 3](#fig3){ref-type="fig"}F).Figure 3Human Skeletal Muscle Mitochondrial Bioenergetics Remain Unaltered with NR Supplementation(A) Mitochondrial respiration of permeabilized muscle fibers upon the addition of complex I and complex II substrates at baseline and after 3 weeks of supplementation of NR and the placebo. MG, malate and glutamate; D, ADP; S, succinate; c, cytochrome C; F, FCCP; Rot, rotenone. Data are normalized to muscle fiber weight.(B) Mitochondrial respiration as per (A), but with the prior addition of the fatty acid conjugate octanoyl-carnitine to malate (MOct).(C) Citrate synthase (CS) activity in human skeletal muscle at baseline and after NR and the placebo.(D) Relative PCR expression of mitochondrial DNA (mtDNA) to nuclear DNA (nDNA) at baseline and after NR and the placebo, expressed as arbitrary units.(E) Western blot showing the expression of selected mitochondrial proteins in skeletal muscle lysates compared to β-actin as housekeeping protein.(F) Western blot showing the expression of acetylation proteins in skeletal muscle lysates compared to β-actin as housekeeping protein.Data are obtained from 12 participants at each phase and wherever relevant are presented as mean ± SEM. Significance was set at p \< 0.05. The absence of significance symbols indicates a lack of statistical significance.

Data from rodents suggest that NAD^+^ supplementation can improve the physiological function in skeletal muscle decline ([@bib9], [@bib25], [@bib50]); thus, we used hand-grip strength as a surrogate marker for muscle function, but one cannot expect hand-grip strength to change after 3 weeks of NR supplementation and without muscle training. Hand-grip strength correlates with leg strength and is used for the diagnosis of sarcopenia and frailty, and it is a better predictor for clinical outcomes than low muscle mass ([@bib41]). A decline in hand-grip strength is observed after the third decade of life (when median peak strength is 51 kg of force in men) ([@bib19]), dropping to median of 33.8 kg of force in our participants. A grip strength of \<30 kg of force in men is a diagnostic criterion for sarcopenia ([@bib15], [@bib16]). After 3 weeks of supplementation, we did not observe any differences in the participants' peak hand-grip strengths (NR 32.5 kg versus placebo 34.7 kg; p = 0.96) or body-weight-adjusted relative strength (NR 2.4 versus placebo 2.3; p = 0.96) between NR and the placebo ([Figure S4](#mmc1){ref-type="supplementary-material"}).

Oral NR Does Not Alter Skeletal Muscle Blood Flow or Substrate Utilization {#sec2.5}
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Recent mouse data showed that NMN increases angiogenesis and muscle blood flow ([@bib17]). Therefore, we used venous occlusive plethysmography to test forearm muscle blood flow in the participants in a non-invasive manner ([@bib30]). At fasting, no NR-mediated differences were detected in muscle blood. Following oral glucose load, muscle blood flow gradually increases, but again with no differences between NR and the placebo ([Figure 4](#fig4){ref-type="fig"}A).Figure 4Forearm Muscle Blood Flow and Substrate Utilization Are Unaffected by NR Supplementation(A) Muscle blood flow using venous occlusive plethysmography at baseline and after the NR and placebo phases. The green dotted line represents when 75 g of oral glucose load was taken.(B and C) Muscle O~2~ consumption (B) and CO~2~ production (C) at baseline and after NR and the placebo. The green dotted line represents when 75 g of oral glucose load was taken.(D and E) Muscle glucose uptake (D) and lactate release (E) at baseline and after NR and the placebo. The green dotted line represents when 75 g of oral glucose load was taken.Data are obtained from 12 participants at each phase and presented as mean ± SEM. Significance was set at p \< 0.05 using a paired t test. The absence of significance symbols indicates a lack of statistical significance.

We then used the arteriovenous difference method (see [Method Details](#sec4.4){ref-type="sec"}) to compare substrate utilization across the forearm muscle (between arterial blood supplying the muscle and venous blood drained from the muscle), with muscle blood flow taken into consideration ([@bib4]). No differences were detected in O~2~ consumption ([Figure 4](#fig4){ref-type="fig"}B) and CO~2~ production ([Figure 4](#fig4){ref-type="fig"}C) between NR and the placebo at the fasting state and in response to oral glucose. Muscle glucose uptake was increased following oral glucose before a gradual decline. No changes were observed in muscle glucose handling with or without NR ([Figure 4](#fig4){ref-type="fig"}D). Oral glucose reduced lactate production from muscle, again without a difference in response between NR and the placebo ([Figure 4](#fig4){ref-type="fig"}E). These data suggest that the skeletal muscle transcriptomic signature of downregulated mitochondrial and glycolysis genes is undetectable when considered at a functional level.

Oral NR Does Not Alter Systemic Cardiometabolic Parameters {#sec2.6}
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Several preclinical studies have described that NAD^+^ supplementation promotes a resistance to weight gain, ameliorates markers of cardiometabolic risk, and improves metabolic flexibility ([@bib71]). As NR increased the circulating levels of the NAD^+^ metabolome, we reasoned that there was increased NAD^+^ availability and turnover in central and peripheral tissues and assessed for resultant cardiometabolic adaptations. Two studies---one of 12 weeks of NR supplementation at 2 g/day in subjects with obesity ([@bib20]) and one of 6 weeks of NR supplementation at 1 g/day in older adults ([@bib46])---suggested potential benefits with respect to fatty liver and blood pressure, respectively. Data for participants at baseline and following NR or the placebo are reported in [Table S1](#mmc1){ref-type="supplementary-material"}. There were no changes in body weight, blood pressure, lipid profile, fasting glucose and insulin ([Table S1](#mmc1){ref-type="supplementary-material"}), and homeostatic model assessment of insulin resistance (HOMA-IR) ([Figure 5](#fig5){ref-type="fig"}A). A rebound increase in non-esterified fatty acids (NEFAs) has previously been associated with the nicotinic acid analog, acipimox ([@bib65]); however, NR did not produce this effect in our trial ([Figure 5](#fig5){ref-type="fig"}B). Glucose handling was studied using an oral glucose tolerance test, with no effect of NR measured in glucose levels during the 2-h test ([Figure 5](#fig5){ref-type="fig"}C). Following the oral glucose load and the consequent insulin stimulation, NEFA levels were appropriately suppressed, and no difference in this response was observed between NR and the placebo ([Figure 5](#fig5){ref-type="fig"}D). We also assessed metabolic flexibility using indirect calorimetry to derive respiratory exchange ratios (RERs; calculated as VCO~2~ expired/VO~2~ consumed), reflecting whole-body metabolic substrate use. Measurements were initiated in the fasted state and monitored during the response to the oral glucose load. The median fasting RER was appropriate at 0.72 and 0.73 for the NR and placebo periods, respectively (p = 0.68). In response to glucose, RER values significantly increased, indicating adequate switching from lipids toward carbohydrate utilization, with no differences in response to 3 weeks of NR supplementation observed at 2 h (RERs 0.83 and 0.84 for NR and the placebo, respectively) ([Figure 5](#fig5){ref-type="fig"}E).Figure 5Systemic Readouts of Metabolism Are Unaltered with NR Supplementation(A) HOMA-IR at baseline and after NR and the placebo.(B) Fasting non-esterified fatty acid (NEFA) level at baseline and after NR and the placebo.(C) Plasma glucose response in a glucose tolerance test at baseline and after NR and the placebo. The green dotted line represents when 75 g of oral glucose load was taken.(D) Plasma NEFA response in a glucose tolerance test at baseline and after NR and the placebo. The green dotted line represents when 75 g of oral glucose load was taken.(E) Respiratory exchange ratio (RER) at baseline and after NR and the placebo. The green dotted line indicates when 75 g of oral glucose was taken.Data are obtained from 12 participants at each phase and presented as mean ± SEM. Significance was set at p \< 0.05. The absence of significance symbols indicates a lack of statistical significance.

Oral NR Depresses Circulating Levels of Inflammatory Cytokines {#sec2.7}
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Chronic inflammation appears to be a consistent feature of aging, even in apparently healthy individuals ([@bib59]), and may contribute to age-related disturbances in metabolic homeostasis ([@bib35]). We hypothesized that NR supplementation would reduce the levels of circulating inflammatory cytokines. We measured multiple inflammatory cytokines (see [Method Details](#sec4.4){ref-type="sec"}), 10 of which were within the assay detection range ([Figure 6](#fig6){ref-type="fig"}). NR significantly decreased the levels of the interleukins IL-6 ([Figure 6](#fig6){ref-type="fig"}A), IL-5 ([Figure 6](#fig6){ref-type="fig"}B), and IL-2 ([Figure 6](#fig6){ref-type="fig"}C) and tumor necrosis factor alpha (TNF-α) ([Figure 6](#fig6){ref-type="fig"}D), compared to baseline. We detected a statistically significant difference in the levels of IL-2 between baseline and the placebo ([Figure 6](#fig6){ref-type="fig"}C) and a lack of a difference in levels of TNF-α between NR and the placebo, despite a difference between NR and baseline ([Figure 6](#fig6){ref-type="fig"}D). This is seemingly due to the NR carry-over effect beyond the washout period, as evident by the period effect analysis ([Figures S5](#mmc1){ref-type="supplementary-material"}A--S5D), confirming that the cohort randomized to the placebo first had no difference in IL-2 between baseline and the placebo ([Figure S5](#mmc1){ref-type="supplementary-material"}C), and there was a difference in TNF-α between NR and the placebo ([Figure S5](#mmc1){ref-type="supplementary-material"}D). No NR-mediated changes were detected in the serum levels of IL-12 ([Figure 6](#fig6){ref-type="fig"}E), IL-8 ([Figure 6](#fig6){ref-type="fig"}F), interferon-gamma (IFN-g) ([Figure 6](#fig6){ref-type="fig"}G), monocyte chemoattractant protein-1 (MCP-1) ([Figure 6](#fig6){ref-type="fig"}H), macrophage inflammatory protein-1 beta (MIP-1B) ([Figure 6](#fig6){ref-type="fig"}I), and high-sensitivity C-reactive protein (hsCRP) ([Figure 6](#fig6){ref-type="fig"}J). Thus, it will be interesting to further investigate depressed IL-6, IL-5, IL-2, and TNF-α as biomarkers and/or mediators of oral NR in rodent models and humans.Figure 6NR Supplementation Suppresses the Circulating Levels of Inflammatory Cytokines(A--J) Levels of serum inflammatory cytokines at baseline and after each of the NR and placebo phases, including (A) interleukin 6 (IL-6), (B) interleukin 5 (IL-5), (C) interleukin 2 (IL-2), (D) tumor necrosis factor alpha (TNF-α), (E) interleukin 12 (IL-12), (F) interleukin 8 (IL-8), (G) interferon-gamma (IFN-g), (H) monocyte chemoattractant protein-1 (MCP-1), (I) macrophage inflammatory protein-1 beta (MIP-1B), and (J) high-sensitivity C-reactive protein (hsCRP). Data are obtained from 12 participants at each phase and presented as mean ± SEM. Significance was set at p \< 0.05 using paired t test. The absence of significance symbols indicates a lack of statistical significance.

Discussion {#sec3}
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The NAD^+^ precursor NR has been studied extensively in animal and cell models. Its application *in vivo* has produced impressive results ameliorating metabolic dysfunction and muscle decline ([@bib23]). No data exist on whether oral NR is available to human skeletal muscle, and data on tissue NAD^+^ content during aging are sparse, as are the consequences of NR supplementation in aged humans.

Using a robust clinical trial design, we show that 21 days of NR supplementation is safe and well tolerated in an aged male cohort and leads to an augmented NAD^+^ metabolome in whole blood, corroborating data recently reported by others ([@bib46], [@bib63]). The median BMI in this trial is 26.6 kg/m^2^ (i.e., slightly overweight), but this is highly prevalent in aged populations ([@bib68]) and may not indicate an unhealthy state ([@bib56]).

Experiments in genetic mouse models have shown that oral NR is available to cardiac ([@bib18]) and skeletal ([@bib25]) muscle, though it was also suggested that the benefit of extrahepatic NAD^+^ from oral NR is mediated by circulating NAM ([@bib45]). Here, we show that oral NR increased human skeletal muscle NAAD, which was previously reported as a more sensitive marker of increased NAD^+^ metabolism than NAD^+^ per se ([@bib63]), as well as MeNAM, Me-4-py, and Me-2-pywithout a rise in circulating NAM. Previous preclinical studies have established that oral NR is able to functionally restore muscle NAD^+^ despite a loss of NAM salvage ([@bib25], [@bib18]). Although it is clear in rodent models that NR requires NR kinase activity in muscle ([@bib57], [@bib24]), further studies are required to understand NR dynamics in human muscle cells and tissues. Increased circulating levels of MeNAM and expression of its generating enzyme nicotinamide-N-methyltransferase (NNMT) have been associated with insulin resistance and type 2 diabetes ([@bib37], [@bib44]). However, the NR-mediated abundance of MeNAM did not alter glucose tolerance or substrate utilization in our study.

The levels of elevated NAM excretory products in skeletal muscle may be a result of pre-existing NAD^+^ sufficiency in this aged cohort and may explain the lack of effect on mitochondrial, physiological, and cardiometabolic parameters. A limited number of studies have reported minor age-related declines in NAD^+^ in human tissues ([@bib47], [@bib10], [@bib73], [@bib11]), but these data are non-conclusive. It is likely that a "second hit" arises during chronological aging that leads to tissue NAD^+^ decline and predisposes to age-related disease and frailty. This "second hit" may be conditions of metabolic stresses such as physical inactivity, chronic inflammation, or presence of a pre-existing cardiometabolic disease (e.g., obesity), and it may implicate downregulated NAMPT ([@bib13], [@bib35]), depressed hepatic NADP(H) ([@bib64]), and/or activation of CD38 ([@bib8], [@bib14]). Clearly, more human data are needed to delineate the relationship between aging and NAD^+^ metabolism.

We note a median hand-grip strength in our participants of 33.8 kg of force, consistent with muscle aging for men in their eighth decade and likely associated impairment in mitochondrial function. Our data suggest that 3 weeks of NR supplementation without concomitant muscle training is insufficient for increased strength.

NAD+ supplementation studies in rodents showed positive effects on muscle structural proteins ([@bib25], [@bib58], [@bib72]). It has been suggested that for muscle cell membranes, there is a capacity for NAD^+^-mediated ADP-ribosylation of integrin receptors that augment integrin and laminin binding and mobilize paxillin to bind adhesion complexes ([@bib29], [@bib28]). Differential gene expression analysis may support this link and may highlight a potential role for NAD^+^ in the maintenance of skeletal muscle architecture, although this NR-induced transcriptomic signature appears to have no functional consequences at the protein level after the 21-day supplementation period. This observation may be important as we consider defective integrin and laminin structures such as in the context of muscular dystrophies ([@bib48], [@bib49]). Our data suggest a downregulation of gene sets associated with glycolysis and mitochondrial function, yet our measurements of mitochondrial respiration, citrate synthase activity, and mitochondrial copy number were unaltered. Again, expression levels of proteins involved in glycolytic and mitochondrial metabolism were unchanged with NR in this study. The downregulation of energy-generating processes may be reminiscent of mechanisms associated with calorie restriction ([@bib31], [@bib36], [@bib43]) or increased mitochondrial quality control, as has been observed in blood stem cells ([@bib66]), or it may suggest that NR can "tune" the expression of energy metabolism pathways to permit a more efficient and potentially stress-resilient mitochondrial environment. It will be interesting to further investigate those transcriptional changes in cell culture and animal models.

Some preclinical studies have reported that NR reduced macrophage infiltration in damaged muscle ([@bib58], [@bib72]) and attenuated plasma TNF-α in models of fatty liver disease ([@bib26]). We show significant suppression of a number of circulating inflammatory cytokines. Studies are needed to explore the underlying mechanisms that mediate these NR-mediated anti-inflammatory effects. Of note, the expression of the NAD^+^-consuming enzyme CD38 increases in inflammatory cells with inflammation ([@bib2]), as well as in the blood of aged humans ([@bib55]). Supplementing NAD^+^ in this context may be a mechanism mediating the NR-induced anti-inflammatory effects. Though chronic inflammation is a hallmark feature of aging ([@bib59]), use of NR may yet find utility in other chronic inflammatory disorders, such as chronic obstructive pulmonary disease or rheumatoid arthritis, and is worthy of further investigation.

Conclusions {#sec3.1}
-----------

We report that oral NR augments the aged human skeletal muscle NAD^+^ metabolome while inducing a transcriptional signature without affecting mitochondrial function or systemic cardiometabolic parameters. The targeted NAD^+^ metabolome analysis suggests pre-existing NAD^+^ sufficiency, despite hand-grip strength consistent with muscle aging. Our data may suggest that chronological age per se may not be a major factor in altering muscle and brain NAD^+^ metabolism, unlike aged laboratory mice. A limitation of this trial may be the number of participants or the duration of NR administration; however, the sample size was sufficient to detect NR-driven changes in the NAD^+^ metabolome, muscle transcriptional signature, and inflammatory profile. The transcriptional downregulation of mitochondrial gene sets also argues against the lack of a bioenergetic NR effect being due to the sample size. Further studies are needed to conceptualize some of the NR-mediated changes in this experimental medicine study.

Overall, these studies support that oral NR is available to human skeletal muscle, and they reveal anti-inflammatory NR properties, both of which may be beneficial in the context of aging, muscle, or inflammatory disease groups.

STAR★Methods {#sec4}
============

Key Resources Table {#sec4.1}
-------------------

REAGENT or RESOURCESOURCEIDENTIFIER**Antibodies**Custom made secondary antibodies (rabbit & mouse)MRC Phosphorylation and Ubiquitylation unit, University of DundeeN/Aα-Acetyl lysineCytoskeletonCat\#AAC03α-Alpha tubulinSanta-CruzCat\#sc-8035α-ANXA1SigmaCat\#HPA011272α-Beta actinSigmaCat\#A5441α-OXPHOSAbcamCat\#ab110413α-PGK1AbcamCat\#ab199438α-PGM1Protein TechCat\#15161-1-APα-PKM1Protein TechCat\#15821-1-AP**Chemicals, Peptides, and Recombinant Proteins**^13^C labeled nucleotides, nucleosides[@bib62]N/A^18^O- nicotinamide[@bib39]N/A^18^O nicotinamide riboside[@bib70]N/A2-mercaptoethanolVWRCat\#441435CAcetonitrile, Optima LC/MSFisher ScientificCat\#A955-4Adenosine 5′-triphosphate disodium salt hydrateSigma-AldrichCat\#A2383Adenosine 5′diphosphateSigma-AldrichCat\#A5285Adenosine diphosphateSigma-AldrichCat\#1905Adenosine monophosphateSigma-AldrichCat\#A2252ADPRSigma-AldrichCat\#A0752Ammonium acetateSigma-AldrichCat\#238074Antimycin ASigma-AldrichCat\#A8674BenazmidineMilliporeCat\#S7124222Bovine serum albuminSigmaCat\#A2153BSA, essentially fatty acid freeSigma-AldrichCat\#A6003Calcium carbonateSigma-AldrichCat\#C4830Catalase from bovine liverSigma-AldrichCat\#C9322Chemiluminescent HRP substrateMilliporeCat\#WBKLS0500CytidineSigma-AldrichCat\#C122106Cytochrome *c*Sigma-AldrichCat\#C7752d~3~, ^18^O methyl nicotinamide[@bib63]N/Ad~4~-nicotinic acidCDN isotopesCat\#D-4368DithiotreitolSigma-AldrichCat\#D0632DL-octanoyl carnitine-HClTocris bioscienceCat\#605Dnase I, Rnase-freeThermo ScientificCat\#EN0521EDTASigmaCat\#E1644EGTASigma-AldrichCat\#E4378EGTASigmaCat\#E4378FCCPSigma-AldrichCat\#2920Glucose reagentWerfen LtdCat\#00018250740Glutamic acid monosodium salt hydrateSigma-AldrichCat\#G1626Glycerol reagentRandoxCat\#GY105GlycineVWRCat\#0167HEPESFlukaCat\#BP310-1ImidazoleSigma-AldrichCat\#56750InosineSigma-AldrichCat\#I4125Inosine monophosphateSigma-AldrichCat\#57510Lactate ReagentRandoxCat\#LC2389Lactobionic acidSigma-AldrichCat\#153516LDS sample bufferInvitrogenCat\#NP0008Magnesium chloride hexahydrateSigma-AldrichCat\#M2670Malic acidSigma-AldrichCat\#M1000MES Free Acid HydrateSigma-AldrichCat\#M8250N1-methyl-2-pyridone-5-carboxamideTLC Pharmaceutical StandardsCat\#N-0621N1-methyl-4-pyridone-3-carboxamideTLC Pharmaceutical StandardsCat\#N-0627NAADSigma-AldrichCat\#N4256NADSigma-AldrichCat\#N0632NADPSigma-AldrichCat\#N5755NARgiftN/AN-d~3~ methyl −4 pyridone-3-carboxamideTLC Pharmaceutical StandardsCat\#N-0628NEFA reagentRandoxCat\#FA115NicotinamideSigma-AldrichCat\#72340nicotinamide N-oxideSigma-AldrichCat\#N3258Nicotinamide ribosideChromaDexCat\#ASB-00014315Nicotinic acidSigma-AldrichCat\#N4126N-methyl nicotinamideCayman ChemicalCat\#16604NMNSigma-AldrichCat\#N3501Phosphocreatine disodium salt hydrateSigma-AldrichCat\#P7936Potassium phosphate monobasicSigma-AldrichCat\#P9791Protease inhibitor cocktailRocheCat\#11873580001RotenoneSigma-AldrichCat\#R8875Saponin from Quillaja barkSigma-AldrichCat\#S7900SDSITWCat\#A1112Skimmed milk powderCell SignalingCat\#9999Sodium chlorideVWRCat\#27800.360Sodium fluorideAlfa AesarCat\#7681-49-4Sodium orthovanodateAldrichCat\#450243Sodium pyrophosphateSigmaCat\#221368Sodium succinate dibasic hexahydrateSigma-AldrichCat\#S2378SucroseSigma-AldrichCat\#S9378SucroseVWRCat\#0335TaurineSigma-AldrichCat\#T8691TRI ReagentSigma-AldrichCat\#T9414Triglyceride reagentWerfen LtdCat\#00018258740Tris-baseFisherCat\#BP152Tris-HCLFisherCat\#BP153Triton X-100SigmaCat\#101634725Tween-20VWRCat\#0777U-^13^C~6~ glucoseCambridge IsotopeCat\#CLM-1396-pkWater, Optima LC/MSFisher ScientificCat\#W6-4β-GlycerophosphateSigmaCat\#G9422**Critical Commercial Assays**Citrate synthase assay kitSigma-AldrichCat\#CS0720Coomassie protein assay reagentThermo FisherCat\#1856209Insulin assayMercodiaCat\#10-1113-01**Deposited Data**Raw and processed data files for RNA sequencingThis paperGEO: [GSE133261](ncbi-geo:GSE133261){#intref0010}**Oligonucleotides**Please refer to [Table S6](#mmc5){ref-type="supplementary-material"}This paperN/A**Other**Precast gelsBioRadCat\#5671084Thermo hypercarb 2.1 × 100 mm column, 3μmFisherCat\#35003-102130

Lead Contact and Materials Availability {#sec4.2}
---------------------------------------

Further information and requests for resources and reagents should be directed to and will be fulfilled by the Lead Contact, Gareth Lavery (<g.g.lavery@bham.ac.uk>). This study did not generate new unique reagents.

Experimental Model and Subject Details {#sec4.3}
--------------------------------------

### Study conduct {#sec4.3.1}

The study was conducted between July 2016 and August 2017 at the National Institute for Health Research/Wellcome Trust Clinical Research Facility at the Queen Elizabeth Hospital Birmingham, UK. The Solihull NRES Committee gave ethical approval (REC reference number 16/WM/0159). All participants provided written informed consent. The study was registered on [www.clinicaltrials.gov](http://www.clinicaltrials.gov){#intref0020} (Identifier: NCT02950441).The study was undertaken according to the principles of the Declaration of Helsinki and followed the Guidelines for Good Clinical Practice.

### Study population {#sec4.3.2}

Aged volunteers were recruited from the Birmingham 1000 Elders group (<https://www.birmingham.ac.uk/research/activity/mds/centres/healthy-ageing/elders.aspx>). All participants fulfilled the inclusion criteria including: male sex, age 70 -- 80 years, BMI 20 -- 30 kg/m^2^, able to discontinue aspirin for 3 days prior to the muscle biopsy, and able to discontinue statins and vitamin D supplements for a week before the study and for the duration of the study. Exclusion criteria included: serious active medical conditions including inflammatory diseases or malignancies, significant past medical history including diabetes mellitus, ischemic heart disease, cerebrovascular disease, respiratory disease requiring medication, or epilepsy, blood pressure \>160/100mmHg, or treatment with oral anti-coagulants.

### Study design {#sec4.3.3}

Single center, double blind, placebo-controlled, and crossover study. Aim was to obtain complete assessments from 12 aged individuals. Participants attended for a screening visit (visit 1) when an informed written consent was obtained after ensuring they fulfil all inclusion criteria. For all subsequent study visits (2 to 5), the participants attended at 08:00 in a fasting state from the night before. Regarding the post-interventions visits (3 and 5), the participants took the last NR/placebo dose 14 h prior to the assessments.

### Randomization and blinding {#sec4.3.4}

Participants were allocated to either NR or placebo. A randomization list was held by the clinical trials pharmacist at the clinical research facility. The study investigators, nurses, and participants were all blinded to the intervention allocation during the trial.

### Intervention {#sec4.3.5}

NR was supplied as 250 mg capsules by the manufacturer (Niagen, ChromaDex, Irvine, CA). Participants received NR 500mg twice daily or matched placebo for 21 days with 21 days washout period between the NR and placebo periods ([Figure S1](#mmc1){ref-type="supplementary-material"}).

### Assessments {#sec4.3.6}

Assessments undertaken in each study visits are detailed in [Figure S1](#mmc1){ref-type="supplementary-material"}.

Method Details {#sec4.4}
--------------

### Blood pressure {#sec4.4.1}

Blood pressure (Welch Allyn, USA) was measured at the start of each visit after an overnight fast. At the trial visits, participants rested for 15mins in a supine position before blood pressure was measured. An appropriately sized cuff was selected to encircle at least 80% of the arm and the same was used every visit for all participants, and on the same arm. Blood pressure was measured in triplicates and the mean was recorded.

### Hand-held dynamometry {#sec4.4.2}

Peak absolute strength (kilograms) and relative handgrip strength (kilograms of force per kilogram of body weight) were measured in triplicate bilaterally using a dynamometer (Takei Instruments, Japan). The highest measurement values were included for analysis.

### Muscle biopsies {#sec4.4.3}

Resting vastus lateralis muscle biopsies were obtained from 12 men by a single investigator (Y.SE) using a percutaneous Bergstrom technique as previously described ([@bib3]) under local anesthesia (1% lignocaine). The biopsy sample (100 -- 150mg) was immediately dried on clean filter paper and approximately 10mg of tissue was cut and placed on ice cold BIOPS buffer for high-resolution respirometry (see below). The rest of the muscle tissue was immediately snap frozen in liquid nitrogen and stored at −80°C pending analysis.

### Indirect calorimetry {#sec4.4.4}

Participants were allowed to rest for 60 mins after insertion of the arterial and venous catheters. Then they lay supine in a comfortable position while a transparent ventilated canopy was placed over their head. Plastic sheet attached to the hood was placed around the subjects to form a seal. The room temperature, barometric pressure, and humidity were measured by a hygrometer (Oregon Scientific). During the measurement period, participants remained supine, and breathed normally. Respiratory measurements, including resting oxygen consumption (VO~2~) and carbon dioxide production (VCO~2~) and respiratory exchange ratio (VCO~2~/VO~2~), using the mixing chamber mode of the metabolic cart (AEI MOXUS II Metabolic System). Measurements were collected at fasting and then every 30 min for 2 h following a 75 g oral glucose load. Measurements for each period lasted 15 mins so the first and last min could be discarded, and the mean value for the middle 5 min was recorded.

### Arterio-venous difference technique {#sec4.4.5}

An arterial catheter was inserted into a radial artery and a retrograde cannula was inserted into in a deep antecubital vein draining a forearm muscle, on the opposite side of the arterial line. To prevent contamination of the muscle venous blood with the mixed blood from the hand, a wrist cuff was inflated to 200mmHg for 3 mins before sampling. Blood sampling was undertaken simultaneously from the arterial and venous sites at fasting and every 20mins following oral glucose load for 120 min.

### Venous occlusive plethysmography {#sec4.4.6}

Forearm muscle blood flow was measured by venous occlusive plethysmography (Hokanson, USA) ([@bib69]) as previously described ([@bib30]). Blood flow measurements were taken immediately after each blood sampling.

NAD^+^ metabolomics {#sec4.5}
-------------------

Muscle tissue was pulverised and approximately 10 mg was used for each of the acid (A) and basic (B) metabolite extraction. For each sample, internal standard mixtures for each of A and B were prepared. The extraction was undertaken using 0.2 mL of ice-cold LC-MS/MS grade methanol and kept on ice before adding 0.3ml of internal standard made in LC-MS grade water. Samples were sonicated in an acetone water bath (at −4°C) for 20 s, placed back on ice, and then incubated at 85°C with constant shaking at 1050 rpm for 5 min. Samples were then placed on ice for 5 min and centrifuged (16.1k x g, 10 min, 4°C). The supernatant was transferred to clean tubes and dried using a speed vacuum. The dried extract was re-suspended in 30 μl of either LC-MS grade water for acid extract or 10 mM ammonium acetate for alkaline extract and centrifuged (16.1k x g, 3 min, 4°C). The supernatant was carefully transferred to a Waters Polypropylene 0.3 mL plastic screw- top vial. The pellet was then dried using a speed vacuum, pellet was weighed, and later used to normalize data that were finally reported as pmol/mg.

Otherwise, muscle, blood, and urine metabolomics were undertaken as previously described ([@bib62], [@bib63]).

### Blood biochemical analysis {#sec4.5.1}

Blood was drawn from the arterial and venous catheters into heparinised blood tubes. Plasma was rapidly separated by centrifugation at 4°C and was then snap frozen. Plasma glucose, NEFA, and lactate concentrations were measured using commercially available kits on an ILAB 650 Clinical Chemistry Analyzer (Werfen Ltd, UK). Insulin was measured using commercially available assay as per the manufacturer's instructions (Mercodia, Sweden). Homeostasis model assessment of insulin resistance (HOMA-IR) was calculated using the formula \[fasting glucose (mmol/L) × fasting insulin (mU/L)/22.5\].

Lipid profile, urea and electrolytes, and thyroid function tests were all measured on the Roche Modular Platforms (Roche, Switzerland). Full blood count was measured on a Beckman Coulter DxH analyzer (USA).

### High-resolution respirometry on permeabilized muscle fibers {#sec4.5.2}

*Ex vivo* mitochondrial function was determined by measuring oxygen consumption polarographically using a two-chamber Oxygraph-2k (OROBOROS Instruments). Oxygen consumption reflects the first derivative of the oxygen concentration (nmol/ml) in time in the respiration chambers and is termed oxygen flux \[pmol/(s^∗^mg)\], corrected for wet weight muscle tissue (2--5 mg) introduced into the chamber. Measurements were undertaken according to a previously described protocol ([@bib53]). Similar results were obtained if respiration rates were corrected for mitochondrial DNA (mtDNA) copy number or citrate synthase activity.

### Mitochondrial density assessments {#sec4.5.3}

For citrate synthase activity, 5mg of snap frozen human muscle was used and the measurement was undertaken as previously described ([@bib32]). Mitochondrial DNA (mtDNA) copy number was determined using quantitative real time PCR. mtDNA copy number was calculated from the ratio of NADH dehydrogenase subunit 1 (ND1) to lipoprotein lipase (LPL) (mtDNA/nuclear DNA) as previously described ([@bib54]).

### RNA sequencing {#sec4.5.4}

RNA was extracted from frozen muscle tissue using Tri Reagent (Sigma-Aldrich) following manufacturer's instructions. Sequencing libraries were prepared using RNA (RIN \> 7) with the Lexogen Quantseq3 FWD kit. Libraries were sequenced using HiSeq2000 across 4 flowcells generating 75bp long single ended reads (average read depth of 6-10M/sample, which is higher than the 4M reads / sample required for analysis for this type of library). All samples were prepared and sequenced as a single pool. Trimmomatic software (v0.32) and bbduk.sh script (Bbmap suite) were used to trim the ILLUMINA adapters, polyA tails and low quality bases from reads. Trimmed reads were then uniquely aligned to the human genome (hg38) using STAR with default settings (v2.5.2b) and the Gencode (v28, Ensembl release 92) annotation as the reference for splice junctions. Mapped reads were quantified using HT-seq (v0.9.1) using Gencode (v28) genes (-intersection-nonempty flag). Differential gene expression was obtained using DEseq2 with paired baseline and treatment samples.

In this analysis we did not use a cutoff to remove lowly expressed genes. Inclusion of lowly expressed genes (at arbitrary cut-offs) had little bearing on our results (97.8% of differentially expressed genes at p \< 0.05 were identical between no cutoff, and a cut-off of \>3). Of note, volcano plot was drawn with a cut-off (\> 3) in order to visualize the typical "V" shape using R. Differentially expressed genes between baseline (control) and NR treated samples at p value = \< 0.05 were annotated using Biological processes (BP) gene sets with DAVID tool. We obtained similar results using gene annotation tool within Gene Set Enrichment Analysis (GSEA) suite ([@bib61], [@bib42]) for gene sets from KEGG pathways and C5-Biological processes.

In addition, we have used GSEA analysis tool to interrogate specific gene sets against our pre-ranked expression data (Control versus NR treatment). GSEA calculates an Enrichment Score (ES) by scanning a ranked-ordered list of genes (according to significance of differential expression (-log10 p value), increasing a running-sum statistic when a gene is in the gene set and decreasing it when it is not. The top of this list (red) contains genes upregulated upon NR+ treatment while the bottom of the list (blue) represents downregulated genes. Each time a gene from the interrogated gene set (i.e., Glycolysis, mitochondria, TCA cycle) is found along the list, a vertical black bar is plotted (hit). If the "hits" accumulate at the bottom of the list, then this gene set is enriched in upregulated genes (and vice versa). If interrogated genes are distributed homogenously across the rank ordered list of genes, then that gene set is not enriched in any of the gene expression profiles (i.e., control gene sets of similar expression levels to interrogated gene sets). GSEA was used in pre-ranked mode with parameters -norm meandiv -nperm 1000 -scoring_scheme weighted. 10 gene sets of equal size and similar expression levels to the interrogated gene sets were generated using a custom pipeline in R (available upon request). We have interrogated the following gene sets: GO0048870; cell motility, GO0030029; actin filament based process, GO0022610; Biological cell adhesion, (also GO0007155 cell adhesion with similar results), M15112: Wong Mitochondria gene module, M3985: KEGG citrate cycle TCA cycle, merge of M15109: BIOCARTA Glycolysis pathway and M5113: REACTOME glycolysis.

### Protein immunoblotting {#sec4.5.5}

Muscle biopsies were homogenized in ice-cold sucrose lysis buffer (50 mM Tris/HCl (pH 7.5), 250 mM Sucrose, 10mM Na-β-Glycerophosphate, 5mM Na-Pyrophosphate, 1mM Benazmidine, 1 mM EDTA, 1 mM EGTA, 1% Triton X-100, 1 mM Na3VO4, 50 mM NaF, 0.1% β-Mercaptoethanol, supplemented with protease inhibitor cocktail). Samples (40-100μg of protein extract) were loaded into 4%--15% Tris/Glycine precast gels (BioRad) prior to electrophoresis. Proteins were transferred onto PVDF membranes (Millipore) for 1h at 100V. A 5% skimmed milk solution made up with Tris-buffered saline Tween-20 (TBS-T, 0.137M NaCl, 0.02M Tris-base 7.5pH, 0.1% Tween-20) was used to block each membrane for 1h before being incubated overnight at 4°C with appropriate primary antibodies. Membranes were washed in TBS-T three times prior to incubation in horse radish peroxidase-conjugated secondary antibody at room temperature for 1h. Membranes were then washed in TBS-T prior to antibody detection via enhanced chemiluminescence horseradish peroxidase substrate detection kit (Millipore). Images were undertaken using a G:Box Chemi-XR5 (Syngene).

### Inflammatory cytokines {#sec4.5.6}

We performed a multiplex cytokine bead assay using the Bio-Plex Pro Human Cytkine 17-plex panel analyzed with a flow-cytometry based Luminex 200 reader. The levels of IL-1b, IL-2, IL-4, IL-5, IL-6, IL-7, IL-8, IL-10, IL-12, IL-13, IL-17, G-CSF, GM-CSF, IFN-g, MCP-1, MIP-1b, and TNF-a were measured on the participants' sera as per the manufacturer's instructions. Only IL-2, IL-5, IL-6, IL-8, IL-12, IFN-g, TNF-a, MCP-1, and MIP-1b were within detection range. High sensitive CRP was measured using CRPHS: ACN 217 on COBAS 6000 analyzer (Roche, USA). All measurements were undertaken in duplicates.

Quantification and Statistical Analysis {#sec4.6}
---------------------------------------

Sample size for this experimental medicine study was decided upon based on previous experience from studies using the same methodological design, whereby the proposed sample size was sufficient to detect significant differences at the 5% level. The analysis was based on data from all participants who were randomized, and completed all the study visits and assessments. Outcome data were reported as mean ± SEM (or median and quartiles where appropriate). In the NR supplementation study, comparisons of participants between placebo and NR supplementation phases were undertaken using paired t tests. In addition, further data analysis taking into account the period effect was undertaken, by grouping the participants into those who were randomized to NR first and second. This is to look for carryover effect across all analyses. Wherever relevant, area under the curve was calculated using the trapezoid method. Data were analyzed using IBM SPSS Statistics version 22 and GraphPad Prism version 7.0.

Data and Code Availability {#sec4.7}
--------------------------

Raw read files and processed data files for RNA sequencing can be found at the NCBI Gene Expression Omnibus (GEO) database (GSE133261). Scripts and other bioinformatics pipelines used to analyze RNA sequencing data can be found at <https://github.com/iakerman/Quantseq>.

Supplemental Information {#app2}
========================

Document S1. Figures S1--S5 and Tables S1 and S2Table S3. NAD+ Metabolomics Data in Skeletal Muscle, Blood, and Urine after Each of the Baseline, Nicotinamide Riboside (NR), Placebo, and Washout Phases (N = 12), Related to Figure 1Table S4. List of Genes Up- and Downregulated in Skeletal Muscle Biopsies upon NR Supplementation (N = 12), Related to Figure 2Table S5. List of Genes in Gene Ontology Terms Related to Processes Up- and Downregulated in Skeletal Muscle Biopsies upon NR Supplementation (N = 12), Related to Figure 2Table S6. List of Primers Used in the Molecular Analysis, Related to STAR MethodsDocument S2. Article plus Supplemental Information

This work was supported by a Medical Research Council (MRC) Confidence in Concept (CiC) award (A.P. and G.G.L.; CiC4/21); a Clinical Research Fellowship from the MRC Arthritis UK Centre for Musculoskeletal Ageing Research (Y.S.E); a Wellcome Trust Senior Fellowship (G.G.L; 104612/Z/14/Z); a Marie Sklodowska-Curie grant (A.G; 705869); and the Roy J. Carver Trust and NIH (C.B; R01HL147545). We thank ChromaDex (Irvine, California) for providing NR and placebo capsules. We also thank all staff at the NIHR/Wellcome Trust Clinical Research Facility - Queen Elizabeth Hospital Birmingham. We acknowledge Professor Alexandra Sinclair, Professor Jeremy Tomlinson, Dr. Zaki Hassan-Smith, and Dr. Keira Markey for their helpful discussions; Dr. Alpesh Thakker for his input into high-resolution respirometry; and Professor Janet Lord for her help with recruitment of participants.

Author Contributions {#sec5}
====================

Y.S.E. and G.G.L. conceived the trial. Y.S.E., K.N.M., A.P., and G.G.L. designed the trial. Y.S.E. conducted the clinical trial. Y.S.E., K.K., D.A.T., and A.G. conducted and supported the high-resolution mitochondrial respirometry. I.A. performed bioinformatic and molecular pathway analyses. Y.S.E., C.L.D., D.M.C., L.O., A.S., and Y.-C.L. contributed to molecular analysis. G.A.W. contributed helpful discussions and supported blood analyses. R.S.F., M.S.S., and C.B. undertook targeted metabolomics and quantitation of NAD^+^-related metabolites, while Y.S.E. analyzed the data. C.V.B., N.J., M.W., and S.J.E.L. supported aspects of the *in vivo* studies. Y.S.E. and P.N. conducted statistical analyses. Y.S.E., C.B., and G.G.L. took the leading roles in writing the manuscript and creating the figures. All authors contributed to the editing and proofreading of the final draft.

Declaration of Interests {#sec6}
========================

C.B. is the inventor of patents licensed by ChromaDex, owns stock in ChromaDex, and serves as an adviser to ChromaDex and Cytokinetics. The remaining authors declare no competing interests.

Supplemental Information can be found online at <https://doi.org/10.1016/j.celrep.2019.07.043>.

[^1]: Lead Contact
