Abstract

   Mitochondrial dysfunction and low nicotinamide adenine dinucleotide
   (NAD^+) levels are hallmarks of skeletal muscle ageing and
   sarcopenia^[92]1–[93]3, but it is unclear whether these defects result
   from local changes or can be mediated by systemic or dietary cues. Here
   we report a functional link between circulating levels of the natural
   alkaloid trigonelline, which is structurally related to nicotinic
   acid^[94]4, NAD^+ levels and muscle health in multiple species. In
   humans, serum trigonelline levels are reduced with sarcopenia and
   correlate positively with muscle strength and mitochondrial oxidative
   phosphorylation in skeletal muscle. Using naturally occurring and
   isotopically labelled trigonelline, we demonstrate that trigonelline
   incorporates into the NAD^+ pool and increases NAD^+ levels in
   Caenorhabditis elegans, mice and primary myotubes from healthy
   individuals and individuals with sarcopenia. Mechanistically,
   trigonelline does not activate GPR109A but is metabolized via the
   nicotinate phosphoribosyltransferase/Preiss–Handler pathway^[95]5,[96]6
   across models. In C. elegans, trigonelline improves mitochondrial
   respiration and biogenesis, reduces age-related muscle wasting and
   increases lifespan and mobility through an NAD^+-dependent mechanism
   requiring sirtuin. Dietary trigonelline supplementation in male mice
   enhances muscle strength and prevents fatigue during ageing.
   Collectively, we identify nutritional supplementation of trigonelline
   as an NAD^+-boosting strategy with therapeutic potential for
   age-associated muscle decline.

   Subject terms: Ageing, Cell biology, Energy metabolism
     __________________________________________________________________

   Trigonelline is identified as a human metabolite that can be used for
   NAD^+ synthesis and is shown to improve muscle function in aged worms
   and mice.

Main

   Sarcopenia is the functional decline of skeletal muscle during ageing
   which impairs mobility and leads to loss of physical independence and
   disability^[97]7. Clinically, sarcopenia is characterized by the
   pathological decrease of muscle mass, strength and gait
   speed^[98]3,[99]8,[100]9, and arises from myofibre wasting and a
   combination of molecular and cellular hallmarks of ageing that
   collectively impair contraction^[101]10–[102]12. Among these,
   mitochondrial dysfunction has a prominent
   role^[103]1,[104]3,[105]13–[106]17, with decreased mitochondrial
   biogenesis, altered mitochondrial dynamics and proteostasis, and
   reduced mitochondrial respiration and ATP production being established
   drivers of muscle ageing phenotypes^[107]3,[108]18,[109]19. Given that
   intertissue cross-talk controls the availability of metabolic fuels for
   mitochondrial bioenergetics and influences muscle function and quality
   of life, research efforts have also characterized systemic
   contributions to sarcopenia. These include chronic low-grade
   inflammation via pro-inflammatory cytokines, altered metabolic fluxes
   via reduced circulating levels of anabolic amino acids and
   perturbations of glucose, vitamin, and lipid
   metabolism^[110]20–[111]25.

   The Multi-Ethnic Molecular determinants of Sarcopenia (MEMOSA) study
   recently identified mitochondrial dysfunction and declined NAD^+ levels
   as prominent molecular hallmarks of sarcopenia in human cohorts from
   different geographies^[112]3. NAD^+ is an essential coenzyme, derived
   from precursors of the vitamin B[3] family, and a key cofactor for
   cellular and organismal metabolism. Mammals can produce NAD^+ from
   different dietary precursors; these include nicotinamide riboside (NR)
   and nicotinamide mononucleotide (NMN), converted via the nicotinamide
   riboside kinase (NRK) pathway, nicotinic acid (NA), also known as
   niacin, which is metabolized via the nicotinate
   phosphoribosyltransferase (NAPRT)-dependent Preiss–Handler pathway,
   tryptophan used in the de novo pathway for NAD^+ biosynthesis, and
   nicotinamide (NAM), which generates NAD^+ via the nicotinamide
   phosphoribosyltransferase (NAMPT) salvage pathway^[113]26. NAD^+ levels
   decline during ageing in several metabolic tissues in rodents and
   humans^[114]1,[115]17,[116]27,[117]28, including in skeletal muscle
   where there is replicated clinical evidence of age-related NAD^+
   deficiency^[118]2,[119]3. However, it is still largely uncharacterized
   whether alterations in muscle mitochondrial and NAD^+ homeostasis are
   reflected in the circulating metabolome, and thus could be used to
   define clinical biomarkers and therapeutic interventions to manage
   later life muscle decline.

   To complement our previous analyses of muscle biopsies of human
   sarcopenia^[120]3 and understand if mitochondrial dysfunction and
   altered NAD^+ metabolism could be linked to systemic changes, we
   investigated serum levels of the kynurenine / vitamin B metabolome in
   individuals with sarcopenia versus healthy controls from the MEMOSA
   cohort (Extended Data Table [121]1). No changes were observed during
   sarcopenia for most of the metabolites analysed, including the vitamin
   B[3] forms that could act as potential NAD^+ precursors. However,
   patients with sarcopenia had lower circulating concentrations of
   trigonelline, a natural alkaloid found in plants^[122]4 and animals,
   including in humans^[123]29,[124]30 (Fig. [125]1a). Trigonelline levels
   were positively correlated with muscle mass assessed via Appendicular
   Lean Mass Index (ALMI) measured using dual-energy X-ray absorptiometry
   (DXA), grip strength and gait speed, all parameters used in the
   clinical definition of sarcopenia^[126]3,[127]9 (Fig. [128]1b). Serum
   levels of trigonelline are also associated with NAD^+ levels in
   skeletal muscle (Extended Data Fig. [129]1a), which we previously found
   to be linked with clinical measures of sarcopenia and muscle
   health^[130]3. Finally, gene set enrichment analysis identified a
   positive association between serum trigonelline levels and several
   metabolic and signalling pathways in skeletal muscle, with
   mitochondrial oxidative phosphorylation showing the strongest
   association with trigonelline (Fig. [131]1c,d and Extended Data Fig.
   [132]1b). Analysis of the Bushehr elderly health cohort^[133]31
   (Extended Data Table [134]2) demonstrated that serum trigonelline is
   also associated with muscle function in an independent replication
   study (Extended Data Fig. [135]1c). Dietary records indicate that serum
   trigonelline levels are independent of dietary caffeine and vitamin
   B[3]intake in this cohort, but possibly linked to other dietary
   factors, such as folate and fibre intake (Extended Data Fig. [136]1c
   and Extended Data Table [137]3). In addition, correction for dietary
   caffeine and vitamin B[3] intake did not affect the association between
   trigonelline and muscle strength (Extended Data Table [138]4).
   Collectively, our targeted metabolomic profiling of human sarcopenia
   revealed trigonelline as a new metabolite associated with muscle
   function, mitochondrial metabolism and NAD^+.

Extended Data Table 1.

   Profiling of serum metabolites of kynurenine/NAD^+ metabolism in the
   Singapore Sarcopenia Study of MEMOSA

   [139]graphic file with name 42255_2024_997_Tab1_ESM.jpg
   [140]Open in a new tab

   The serum metabolites of kynurenine/NAD^+ were measured in the
   Singapore sarcopenia study of MEMOSA using LC–MS/MS (n = 40).

Fig. 1. Serum trigonelline is reduced in human sarcopenia and is associated
with mitochondrial and NAD^+ metabolism in skeletal muscle.

   [141]Fig. 1
   [142]Open in a new tab

   a, Serum levels of trigonelline in healthy controls (n = 20) and
   individuals with sarcopenia (n = 20) from the MEMOSA SSS (unpaired,
   two-tailed Student’s t-test). b, Association of serum trigonelline
   levels with ALMI, grip strength and gait speed; the Pearson correlation
   coefficient and its P value were calculated on n = 40 serum samples
   from the SSS. c, SSS muscle RNA-seq association with serum trigonelline
   levels. Gene set enrichment ordered according to the significance of
   enrichment with only the top ten gene sets being reported. A false
   discovery rate (FDR) < 10^−^20 was trimmed at FDR = 10^−^20 (n = 39
   muscle samples). d, Enrichment plot for the hallmark oxidative
   phosphorylation gene set from c. e,f, Relative NAD^+ levels in HSMMs
   after treatment with increasing concentrations of trigonelline, in the
   absence (e) or presence (f) of FK866 (one-way analysis of variance
   (ANOVA), mean ± s.e.m, n = 6 biological replicates per group). g,
   Relative NAD^+ levels in human primary myotubes from healthy controls
   and patients with sarcopenia from the Hertfordshire Sarcopenia Study
   Extension (HSSe) cohort treated ex vivo with or without trigonelline
   (unpaired, two-tailed Student’s t-test, mean ± s.e.m, n = 3 biological
   replicates per group). h, Relative NAD^+ levels in primary myotubes
   from aged mice (22 months) treated ex vivo with trigonelline (unpaired,
   two-tailed Student’s t-test, mean ± s.e.m, n = 8 and n = 9 biological
   replicates per group). *P < 0.05, **P < 0.01, ***P < 0.001,
   ****P < 0.0001. For the individual P values, see Fig. 1 in the
   [143]Source data.

   [144]Source data

Extended Data Fig. 1. Trigonelline increases NAD^+ in mammalian cells and its
serum levels correlate with muscle NAD^+ and Oxphos signatures in humans.

   [145]Extended Data Fig. 1
   [146]Open in a new tab

   a, Correlation of serum trigonelline levels with muscle NAD^+, (n = 10
   validation muscle samples remaining from the MEMOSA SSS cohort). b,
   Heatmap showing the top 50 oxidative phosphorylation genes associated
   with serum trigonelline levels. c, Correlation analysis of serum
   trigonelline levels with appendicular lean mass (ALM) index, grip
   strength, dietary caffeine and vitamin B3 intake in a replication
   cohort; Pearson correlation coefficient and its p-value were calculated
   on n = 186 serum samples from the Bushehr elderly health cohort. d,
   Molecular structures of nicotinic acid or niacin (NA), trigonelline
   which differs from NA only for the methyl group, and the ribosylated
   (rib) molecules nicotinamide riboside (NR) and NAD^+. e, NAD^+ levels
   relative to untreated cells measured in HSMM with or without
   trigonelline for 6 hours or 24 hours (unpaired two-tailed Student’s
   t-test, mean ± s.e.m, n = 10 biological replicates per group). f, NAD^+
   levels relative to untreated control measured in HSMM myotubes
   following 24 h NAD^+ precursors treatment at 1 mM (One-way ANOVA;
   mean ± s.e.m, n = 6 biological replicates per group). g, Naprt mRNA
   expression in the indicated cell lines (n = 4–12 biological replicates
   per group). h, NAD^+ levels relative to untreated control measured in
   HepG2, C2C12 and IM-PTECs cells following 24 h NAD^+ precursors
   treatment (One-way ANOVA; mean ± s.e.m, n = 8–12 biological replicates
   per group). i, Stability assessment of trigonelline, NR and NMN added
   at 1 mM in human serum and incubated at 37 °C for the indicated
   durations (Two-way ANOVA; mean ± s.e.m, n = 3 biological replicates per
   group). j, Percentage increase in NAD^+ levels in trigonelline treated
   primary myotubes from healthy control and sarcopenic HSSe subjects
   relative to untreated cells (unpaired two-tailed Student’s t-test,
   mean ± s.e.m, n = 3 biological replicates per group). k, Correlation
   analysis of serum trigonelline levels with SHMT2 mRNA levels in muscle
   biopsies from the SSS cohort; Pearson correlation coefficient and its
   p-value were calculated on n = 39 serum samples. l, Correlation
   analysis of SHMT2 gene expression in muscle biopsies with grip strength
   and appendicular lean mass index in the SSS cohort; Pearson correlation
   coefficient and its p-value were calculated on n = 40 serum samples. P:
   <0.05 (*); < 0.01 (**); < 0.001 (***); < 0.0001 (****); n.s.,
   non-significant. For all the individual p values, see the Extended Data
   Fig. 1 Source file.

   [147]Source data

Extended Data Table 2.

   Clinical characterization of the Bushehr Elderly Health Cohort

   [148]graphic file with name 42255_2024_997_Tab2_ESM.jpg
   [149]Open in a new tab

   Clinical parameters were measured in 186 older men of the Bushehr
   nutritional epidemiology study aged 60 years and older. EWGSOP:
   European Working Group on Sarcopenia in Older People.

Extended Data Table 3.

   Association of dietary intake with serum trigonelline levels in the
   Bushehr Elderly Health Cohort

   [150]graphic file with name 42255_2024_997_Tab3_ESM.jpg
   [151]Open in a new tab

   Dietary intake assessed using 24-h recall food questionnaire in the
   Bushehr Elderly Health Cohort was associated with serum trigonelline
   levels measured with LC–MS/MS and analysed with Spearman rank
   correlation.

Extended Data Table 4.

   Association of grip strength with serum trigonelline levels corrected
   for dietary intake in the Bushehr Elderly Health Cohort

   [152]graphic file with name 42255_2024_997_Tab4_ESM.jpg
   [153]Open in a new tab

   Handgrip strength in the Bushehr Elderly Health Cohort was associated
   with serum trigonelline levels measured with LC–MS/MS and corrected for
   dietary intake assessed with a 24-h recall food questionnaire and
   analysed with a Spearman partial rank correlation.

   Trigonelline is an N-methylated form of NA (Extended Data Fig. [154]1d)
   that is synthesized by several plant species^[155]4 and is also a
   metabolite produced by the gut microbiome and endogenous metabolism in
   humans^[156]29,[157]30. Based on this structural proximity to NA and
   the established link between muscle NAD^+ and mitochondria in ageing
   and sarcopenia^[158]1,[159]14,[160]15,[161]17, we tested if
   trigonelline could act as an NAD^+ precursor, and directly impact
   NAD^+, mitochondrial and muscle homeostasis. Primary human skeletal
   muscle myotubes (HSMMs) were treated with increasing therapeutic doses
   of trigonelline in the presence or absence of the NAMPT inhibitor FK866
   (refs. ^[162]14,[163]32) to block NAD^+ salvage (Fig. [164]1e,f), thus
   mimicking low NAD^+ via decreased NAMPT expression in human sarcopenic
   muscle^[165]3. Trigonelline increased NAD^+ in control cells (half
   maximal effective concentration (EC[50]) = 315 µM) (Fig. [166]1e),
   fully rescued NAD^+ deficiency in FK866-treated cells (EC[50] = 110 µM)
   (Fig. [167]1f) and also increased NAD^+ levels after prolonged
   treatment (Extended Data Fig. [168]1e). When compared to other NAD^+
   precursors in primary human muscle cells, trigonelline and the salvage
   precursor NAM induced NAD^+ by approximately 50% while the NRK pathway
   precursors NR and NMN^[169]33 increased NAD^+ levels approximately
   twofold (Extended Data Fig. [170]1f). The Preiss–Handler pathway of
   NAD^+ biosynthesis requires the conversion of its substrate NA into NA
   mononucleotide (NAMN) via the rate-limiting enzyme NAPRT^[171]6. When
   we tested trigonelline and other precursors in additional cell lines of
   muscle, liver and kidney, trigonelline or NA failed to raise NAD^+
   levels in HepG2 (ref. ^[172]34) and C2C12 cells, which have low NAPRT
   expression (Extended Data Fig. [173]1g,h), while all precursors had
   similar efficacy in renal proximal tubular epithelial cells (PTECs)
   (Extended Data Fig. [174]1h). To further understand the biological
   activity of these precursors after in vivo absorption, we compared
   their stability in human serum. Trigonelline was remarkably stable in
   serum over 72 h, whereas NR and NMN rapidly disappeared within hours
   after conversion to NAM (Extended Data Fig. [175]1i). Given its lower
   serum levels in individuals with sarcopenia (Fig. [176]1a), we tested
   trigonelline in sarcopenic muscle cells. Treating primary myotubes from
   different donors with sarcopenia and aged-matched healthy
   controls^[177]35 raised cellular NAD^+ (Fig. [178]1g) with comparable
   efficiency between groups (Extended Data Fig. [179]1j). Similarly,
   trigonelline also increased NAD^+ levels in primary myotubes derived
   from aged mice (Fig. [180]1h).

   Trigonelline is N-methylated on its pyridine ring and needs to be
   demethylated before entry into the Preiss–Handler pathway and pyridine
   N-ribosylated for NAD^+ biosynthesis (Extended Data Fig. [181]1d).
   Because there is no trigonelline demethylase identified in plants or in
   mammals^[182]36, we explored potential candidates that could be linked
   to trigonelline demethylation by correlating serum trigonelline levels
   with the expression of genes from human sarcopenic muscle filtered
   using RNA sequencing (RNA-seq) for demethylating or methyltransferase
   activity (Extended Data Table [183]5). SHMT2, which is involved in
   mitochondrial one-carbon metabolism, had the strongest association with
   serum trigonelline (Extended Data Fig. [184]1k), and correlated
   positively with grip strength and muscle mass in humans (Extended Data
   Fig. [185]1l). While this observation uncovers a potential link between
   trigonelline and a methyltransferase involved in one-carbon metabolism
   that will require further exploration, SHMT2 is unlikely to directly
   demethylate trigonelline because it is not a N-methyltransferase and is
   known to cross-talk with NAD^+ metabolism indirectly^[186]37,[187]38.

Extended Data Table 5.

   Association of serum trigonelline levels with muscle mRNA expression of
   demethylase and methyltransferase genes in the Singapore Sarcopenia
   Study

   graphic file with name 42255_2024_997_Tab5_ESM.jpg
   [188]Open in a new tab

   Serum trigonelline levels measured with LC–MS/MS in the Singapore
   Sarcopenia study of MEMOSA was associated with gene expression measured
   using RNA sequencing in muscle biopsies, analysed with a Spearman rank
   correlation, filtered for genes expressed in skeletal muscle and
   annotated with methyltransferase or demethylase activity.

   To assess whether trigonelline is incorporated into the NAD^+ molecule,
   we used an isotopically labelled form of trigonelline carrying a ^13C
   on the carboxylic acid group and three deuterium (^2H) atoms on the
   methyl group (-CD[3]) (Fig. [189]2a). Administration of isotopically
   labelled trigonelline in mice was highly bioavailable with high levels
   of the parent trigonelline molecule detected in the liver,
   gastrocnemius muscle, kidney, blood and urine 2 h after oral intake,
   and were largely cleared after an overnight wash-out (Extended Data
   Fig. [190]2a). This was mirrored by increased NAD^+ content in the
   liver, muscle, kidney and whole blood, detected with either liquid
   chromatography coupled high-resolution mass spectrometry (LC–HRMS)
   (Fig. [191]2b) or an NAD enzymatic assay (Extended Data Fig. [192]2b).
   Based on our labelling strategy, direct incorporation of trigonelline
   into NAD^+ implies the loss of the deuterated methyl group, leading to
   a ^13C-labelled NAD^+ structure with a mass of M + 1 (Fig. [193]2a).
   Treatment of HSMMs with labelled trigonelline significantly increased
   both total cellular NAD^+ and [^13C]-NAD^+ M + 1 (Fig. [194]2c). In
   vivo, administration of labelled trigonelline also significantly
   enriched M + 1 NAD^+ in the liver and whole blood and, to a smaller
   extent, in the muscle, with hepatic incorporation being faster probably
   because of a first-pass effect (Extended Data Fig. [195]2c).
   Altogether, these results demonstrate that trigonelline is a bona fide
   NAD^+ precursor that is directly incorporated in NAD^+ in cells and
   multiple tissues.

Fig. 2. Trigonelline is an NAD^+ precursor and activates mitochondrial
function via the Preiss–Handler pathway.

   [196]Fig. 2
   [197]Open in a new tab

   a, Experimental design of isotope-labelled trigonelline incorporation
   into NAD^+. b, NAD^+ levels measured using LC–HRMS in the liver,
   gastrocnemius muscle and whole blood 2 h or 16 h (overnight) after
   labelled trigonelline gavage in young mice (one-way ANOVA, n = 4–5 mice
   per group). c, NAD^+ levels measured using LC–HRMS in HSMM (left) and
   relative isotopic enrichment of NAD^+ (right) after 24-h incubation
   with 1 mM labelled trigonelline (unpaired, two-tailed Student’s t-test,
   n = 3 biological replicates per group). d, Representation of the
   Preiss–Handler and salvage pathways of NAD^+ production. e, Relative
   NAD^+ levels in HSMMs 48 h after adenoviral infection with a scrambled
   or NAPRT shRNA (unpaired, two-tailed Student’s t-test, n = 6 biological
   replicates per group). f, Relative NAD^+ levels in HSMMs after 24-h
   trigonelline or NR treatment, with or without 2-OHNA co-treatment
   (one-way ANOVA, n = 12 biological replicates per group). g–j, NAD^+
   metabolites measured using LC–HRMS in HSMMs (NAD^+, g; NAAD, h; NAMN,
   i; NA, j) after 24-h incubation with trigonelline in co-treatment with
   FK866, 2-OHNA or their combination (one-way ANOVA, n = 3 biological
   replicates per group). k, Quantitative PCR (qPCR) mRNA expression of
   Naprt in the liver of wild-type (WT) and Naprt knockout (KO) mice
   (one-way ANOVA, n = 4–6 animals per group). l–n, LC–HRMS measurement of
   NA (l) and NAMN (m) levels in the blood and liver, and of NAAD (n) in
   liver 2 h after trigonelline gavage in WT or Naprt KO mice (one-way
   ANOVA, n = 4–6 animals per group). o, Relative NAD^+ levels in HSMMs
   after 72 h trigonelline or NR treatment, with or without co-treatment
   with FK866, 2-OHNA or their combination (one-way ANOVA, n = 10
   biological replicates per group). p, Mitochondrial membrane potential
   (ΔΨm) measured using JC-1 staining in HSMMs treated as in o (one-way
   ANOVA, n = 16 biological replicates per group). q, Maximum oxygen
   consumption rate (OCR) in HSMMs treated as in o after stimulation with
   3 μM carbonyl cyanide-p-trifluoromethoxyphenylhydrazone (one-way ANOVA,
   n = 9–10 biological replicates per group). All data are expressed as
   the mean ± s.e.m with *P < 0.05, **P < 0.01, ***P < 0.001,
   ****P < 0.0001. NS, not significant. Individual P values are reported
   in Fig. 2 of the [198]Source data. a.u., arbitrary units.

   [199]Source data

Extended Data Fig. 2. Trigonelline and Preiss-Handler pathway metabolomics
and effects on NAD^+ levels and mitochondrial homeostasis.

   [200]Extended Data Fig. 2
   [201]Open in a new tab

   a, LC-HRMS measurements of trigonelline levels in liver, gastrocnemius
   muscles, kidney, whole blood and urine of C57BL/6J mice, collected at
   2 hours (2 h) or overnight (o/n) after labelled trigonelline gavage
   (unpaired two-tailed Student’s t-test compared to control group,
   mean ± s.e.m, n = 5 biological replicates per group, except in urine
   samples where only 2 samples were collected in controls and 4 in
   treated group); LOQ, level of quantification. b, NAD^+ levels relative
   to untreated control mice measured in liver, gastrocnemius and kidney
   tissues from the same groups as in (a) (unpaired two-tailed Student’s
   t-test, mean ± s.e.m, n = 5 biological replicates per group). c,
   LC-HRMS-based relative trigonelline incorporation into NAD^+ in the
   same tissues as in (a), related to the NAD^+ measured in Fig. [202]2b
   (unpaired two-tailed Student’s t-test compared to control group,
   mean ± s.e.m, n = 5 biological replicates per group). d, Naprt mRNA
   levels by qPCR in HSMM cells following 48 h incubation with an
   adenovirus carrying either a scrambled or NAPRT shRNA construct
   (unpaired two-tailed Student’s t-test, mean ± s.e.m, n = 3 biological
   replicates per group). e, NAD^+ levels relative to untreated control
   measured in HSMM myotubes following 24 h incubation with NA in presence
   or absence of the NAPRT shRNA construct (unpaired two-tailed Student’s
   t-test, mean ± s.e.m, n = 6 biological replicates per group). f, NAD^+
   levels relative to untreated control measured in HSMM myotubes
   following 24 h trigonelline or NR treatment, in the presence or absence
   of FK866 (one-way ANOVA; mean ± s.e.m, n = 6 biological replicates per
   group). g, NAD^+ levels relative to untreated control measured in HSMM
   myotubes following 24 h trigonelline or NR treatment, in the presence
   of FK866 and with or without co-treatment with 2-OHNA (one-way ANOVA;
   mean ± s.e.m, n = 10 biological replicates per group). h-i, LC-HRMS
   based measurements of trigonelline and NAM in HSMM myotubes treasted as
   in Fig. [203]2g–j (One-way ANOVA, mean ± s.e.m, n = 3 biological
   replicates per group; in red ****, unpaired two-tailed Student’s t-test
   for comparisons in conditions without trigonelline); AU, abitrary
   units. j, Naprt mRNA expression by qPCR in gastrocnemius muscle of
   wildtype (WT) and Naprt knockout (KO) C57BL/6N mice (One-way ANOVA,
   mean ± s.e.m, n = 4–6 animals per group). k, LC-MS measurements of
   trigonelline levels in liver, plasma and gastrocnemius muscles
   harvested 2 hours post trigonelline gavage in WT or Naprt KO mice
   (unpaired two-tailed Student’s t-test, mean ± s.e.m, n = 4–6 animals
   per group); AU, abitrary units. l-n, LC-MS measurements of NAD^+
   metabolites in (l) liver, (m) blood, (n) gastrocnemius of the same
   groups as in (k) (One-way ANOVA, mean ± s.e.m, n = 4–6 animals per
   group); AU, abitrary units. o, Nampt and Nrk mRNA levels by qPCR in
   liver and muscle of the same groups as in (j) (One-way ANOVA,
   mean ± s.e.m, n = 4–6 animals per group). p, Apoptosis detection via
   annexing staining time course in HSMM myotubes following treatments of
   trigonelline, FK866 or 2-OHNA and their combinations. Staurosporin
   (2.5 µM) is used as a positive control for apoptosis induction (two-way
   ANOVA; mean ± s.e.m, n = 6 biological replicates per group). q,
   Seahorse-based substrate-driven OCR measurement in permeabilized HSMM
   myotubes following 24 h trigonelline treatment, in the presence or
   absence of 2-OHNA, and with a sequence of substrate/inhibitors to
   extract contribution of different complexes (one-way ANOVA;
   mean ± s.e.m, n = 11 biological replicates per group). r, GPR109A
   agonist assay in cells incubated with increasing doses of NA or
   trigonelline (mean ± S.D., n = 4). P: <0.05 (*); < 0.01 (**); < 0.001
   (***); < 0.0001 (****); n.s., non-significant. For all the individual p
   values, see the Extended Data Fig. 2 Source file.

   [204]Source data

   Given the similarity of trigonelline with NA and its ability to
   overcome NAD^+ salvage inhibition (Fig. [205]1e,f), but also its
   inefficacy in low NAPRT-expressing cells (Extended Data Fig.
   [206]1g,h), we tested whether NAPRT would be required for the use of
   trigonelline, and used NR as a reference compound able to generate
   NAD^+ independently of the Preiss–Handler pathway (Fig. [207]2d). Short
   hairpin RNA (shRNA)-mediated knockdown of NAPRT in HSMMs blocked the
   generation of NAD^+ by trigonelline or NA (Fig. [208]2e and Extended
   Data Fig. [209]2d,e). Similarly, the NAPRT inhibitor 2-hydroxynicotinic
   acid (2-OHNA)^[210]39 blocked the conversion of trigonelline to NAD^+
   in HSMMs, but as expected did not impair the NAD^+ boosting effect of
   NR (Fig. [211]2f). Conversely, various levels of cellular NAD^+
   depletion by blocking NAD^+ salvage with FK866 for 24 h were rescued by
   trigonelline (Fig. [212]1f and Extended Data Fig. [213]2f), but not
   when co-treating with 2-OHNA (Extended Data Fig. [214]2g). LC–HRMS
   analysis also confirmed that 2-OHNA blocks increased NAD^+ after
   trigonelline treatment (Fig. [215]2g), without affecting cellular
   trigonelline levels in the treated groups (Extended Data Fig. [216]2h).
   Of note, endogenous levels of trigonelline increased in cells treated
   with 2-OHNA in the absence of trigonelline treatment (Extended Data
   Fig. [217]2h), suggesting that there is an endogenous flux of
   trigonelline through NAPRT in muscle cells. After trigonelline
   treatment, the Preiss–Handler pathway-related metabolites nicotinic
   acid adenine dinucleotide (NAAD) and NAMN downstream of
   NAPRT^[218]6,[219]33 were increased and blocked by NAPRT inhibition
   with 2-OHNA (Fig. [220]2h,i), whereas NA, which is upstream of NAPRT,
   was accumulated (Fig. [221]2j); the salvage pathway metabolite NAM did
   not change (Extended Data Fig. [222]2i). Acute dosing of trigonelline
   in wild-type (WT) and Naprt KO mice^[223]40 (Fig. [224]2k and Extended
   Data Fig. [225]2j) equally increased trigonelline in tissues (Extended
   Data Fig. [226]2k), distal NAD^+ metabolite fluxes in the liver and
   blood (Extended Data Fig. [227]2l,m) and tissue NAD^+ levels (Extended
   Data Fig. [228]2l–n). However, in Naprt KO mice treated with
   trigonelline, NA accumulated in the blood and liver (Fig. [229]2l),
   while NAMN was strongly downregulated (Fig. [230]2m); NAAD induction
   was blunted in the liver (Fig. [231]2n), similarly to what is observed
   in HSMMs (Fig. [232]2h,j). After trigonelline gavage, Nampt and Nrk1
   expression was downregulated in the liver, while Nrk1 and Nrk2 were
   upregulated in muscle in both treatment groups (Extended Data Fig.
   [233]2o). This suggests that trigonelline engages the Preiss–Handler
   pathway during first-pass metabolism and that secondary compensations
   through the NR–NMN–NRK pathway also contribute to NAD^+ in Naprt KOs.
   Altogether, our in vitro and in vivo results indicate that trigonelline
   is metabolized through the Preiss–Handler pathway via NAPRT and
   promotes NAD^+ biosynthesis via both direct and indirect mechanisms.

   To compare the functional relevance of modulating NAD^+ metabolism via
   trigonelline and NR through the NAPRT and salvage routes, we measured
   mitochondrial respiration and membrane potential in HSMMs where low
   NAD^+ was modelled with FK866 for 72 h. Like the 24-h treatment
   (Extended Data Fig. [234]2g), trigonelline boosted NAD^+ and this
   effect was blocked by NAPRT inhibition with 2-OHNA (Fig. [235]2o),
   while no detrimental effect on cell viability was observed (Extended
   Data Fig. [236]2p). Low NAD^+ in response to FK866 lowered the
   mitochondrial membrane potential (Fig. [237]2p), which was fully
   restored by trigonelline and NR, while 2-OHNA abolished the effects of
   trigonelline but not NR (Fig. [238]2p). FK866 treatment also
   significantly decreased maximal mitochondrial respiration (Fig.
   [239]2q), which was also rescued by trigonelline but blocked when
   trigonelline was co-treated with 2-OHNA (Fig. [240]2q). In contrast, NR
   was still able to bypass NAPRT inhibition and increase respiration
   (Fig. [241]2q). Finally, respirometry performed with mitochondrial
   complex inhibitors revealed higher levels of complex II-driven and
   complex IV-driven respiration in trigonelline-treated cells, again
   dependent on NAPRT (Extended Data Fig. [242]2q). The clinical use of NA
   is limited by its effects on skin flushing caused by binding to the G
   protein-coupled receptor 109A (GPR109A)^[243]41. Therefore, we also
   tested whether trigonelline may activate this receptor using a
   GPR109A-overexpressing stable cell line coupled to β-arrestin-based
   detection, and validated the assay with NA (EC[50] = 2.5 µM) (Extended
   Data Fig. [244]2r). Trigonelline did not activate GPR109A when tested
   up to 1 mM, a dose 400-fold higher than the EC[50] of NA (Extended Data
   Fig. [245]2r), suggesting that it could be better tolerated than NA.
   Collectively, these results demonstrate that trigonelline functionally
   rescues NAD^+ deficiency and confirm that trigonelline requires NAPRT
   and a functional Preiss–Handler pathway for physiological activity on
   mitochondrial bioenergetics.

   Lower NAD^+ levels during ageing are linked to mitochondrial
   dysfunction, muscle decline and reduced fitness and lifespan in lower
   organisms, such as nematodes^[246]14,[247]42–[248]45. Given that
   NAD^+-enhancing interventions can improve these
   phenotypes^[249]14,[250]42–[251]45, we next tested the impact of
   trigonelline during ageing in Caenorhabditis elegans by treating N2 WT
   worms starting from day 1 of adulthood (Fig. [252]3a). Trigonelline
   significantly extended the lifespan, with comparable effects to
   equimolar levels of NR (Fig. [253]3b and Extended Data Fig. [254]3a).
   Akin to that observed in human and mouse cells and tissues,
   trigonelline raised NAD^+ levels in aged nematodes (Fig. [255]3c); its
   NAD^+-boosting effects were comparable to NA and NAM, and partially
   lower than NR and NMN (Extended Data Fig. [256]3b). Trigonelline
   increased mitochondrial content assessed using the mitochondrial
   DNA/nuclear DNA ratio (Fig. [257]3d), activated the expression of genes
   linked to mitochondrial respiration and proteostasis (Fig. [258]3e),
   similar to NR (Extended Data Fig. [259]3c and^[260]14,[261]43), and
   increased mitochondrial respiration (Fig. [262]3f and Extended Data
   Fig. [263]3d). We next evaluated muscle structure and mobility in
   ageing nematodes. Trigonelline supplementation improved myofibre
   integrity during ageing (Fig. [264]3g and Extended Data Fig. [265]3e);
   this was mirrored by reduced worm paralysis (Fig. [266]3h) and
   increased spontaneous mobility (Fig. [267]3i). Interestingly, worms
   treated with trigonelline later in adulthood showed only a mild
   lifespan extension (Extended Data Fig. [268]3f) but maintained better
   mobility than control animals during ageing (Extended Data Fig.
   [269]3g), indicating benefits of trigonelline on health span even when
   treating for a shorter duration and later in life. Importantly, the
   effects of trigonelline on NAD^+ and mitochondrial content were lost
   with knockdown of the Naprt worm orthologue nprt-1 (Fig. [270]3j,k), in
   line with what was observed in HSMMs. Knockdown of nprt-1 or sir-2.1
   (Extended Data Fig. [271]3j), the predominant sirtuin in N2 worms
   (Extended Data Fig. [272]3k), both blunted the lifespan-extending
   effects and mobility benefits of trigonelline (Fig. [273]3l–n), in line
   with similar effects observed with NA (Extended Data Fig. [274]3h,i)
   and other NAD^+ boosters^[275]14,[276]42–[277]45. As expected,
   trigonelline increased NAD^+ levels after sir-2.1 knockdown because
   sirtuins act as NAD^+ sensors downstream of NAD^+ biosynthesis and
   sir-2.1 knockdown did not change baseline NAD^+ levels (Extended Data
   Fig. [278]3l). Together, these results validate the physiological
   requirement of the Preiss–Handler pathway for longevity, mitochondrial
   and functional benefits of trigonelline in an in vivo model, and
   demonstrate that the observed benefits on ageing and health span
   require the NAD^+-dependent sirtuin deacetylase.

Fig. 3. Trigonelline supplementation enhances the lifespan and ameliorates
age-related muscle decline and mitochondrial dysfunction in C. elegans.

   [279]Fig. 3
   [280]Open in a new tab

   a, Experimental design of compound treatments at 1 mM started from day
   1 of adulthood (D1) in N2 WT worms. Illustration created with
   [281]BioRender.com. b, Lifespan in trigonelline-treated worms (log-rank
   test, n = 90 worms per group). c, Relative NAD^+ levels in aged worms
   on day 8 (D8). Unpaired, two-tailed Student’s t-test, n = 6 biological
   replicates per group). d, mitochondrial and nuclear DNA in aged worms
   (D8). Unpaired, two-tailed Student’s t-test, n = 12 worms per group).
   e, mRNA expression of mitochondrial genes in worms treated with
   trigonelline (unpaired, two-tailed Student’s t-test, n = 6 biological
   replicates per group). f, Basal and maximal OCR in L4 worms treated
   from the embryo stage (unpaired, two-tailed Student’s t-test, n = 36
   animals per group). g, Confocal images (left) and quantitative
   integrity scoring (right) of green fluorescent protein (GFP)-labelled
   muscle fibres in adult (D1) and aged (D11) RAW1596 (myo-3p::GFP) worms
   (one-way ANOVA, n = 6 worms and 9–31 sarcomeres per group). Scale bar,
   10 µm. h, Percentage of paralyzed aged worms (D11) (n = 3 independent
   experiments, unpaired two-tailed Student’s t-test). i, Spontaneous
   mobility of worms at different ages (unpaired, two-tailed Student’s
   t-test, n = 33–49 worms per group). j, Relative NAD^+ levels in D1
   adult worms treated from the embryo stage and fed with control (empty
   vector (e.v.)) or nrpt-1 RNA interference (RNAi) (one-way ANOVA,
   n = 6–14 biological replicates per group). k, Mitochondrial and nuclear
   DNA in worms treated as in j (one-way ANOVA, n = 10–12 animals per
   group). l,m, Lifespan of control (e.v.), nrpt-1 RNAi (l) and sir-2.1
   RNAi (m) worms (log-rank test, n = 100 animals per group). n,
   Spontaneous mobility of control (e.v.), nrpt-1 RNAi and sir-2.1 RNAi
   worms at D6 (unpaired, two-tailed Student’s t-test, n = 63–123
   individual worms per group). All data are expressed as the
   mean ± s.e.m. with *P < 0.05, **P < 0.01, ***P < 0.001, ****P < 0.0001.
   Individual P values of the worms per group are reported in Fig. 3 of
   the [282]Source data.

   [283]Source data

Extended Data Fig. 3. Effects of the NAD^+ boosters trigonelline, NR and NA
on nematode phenotypes.

   [284]Extended Data Fig. 3
   [285]Open in a new tab

   a, Lifespan assessment in N2 worms treated with or without NR as
   designed in Fig. [286]2a (log-rank test, n = 90 animals per group). b,
   NAD^+ levels (% control) in D1 adult N2 worms treated with NAD^+
   precursors from embryo stage (One-way ANOVA, mean ± s.e.m, n = 6–8
   biological replicates per group). c, mRNA levels by qPCR of genes
   regulating mitochondrial function in N2 worms treated with NR from Day
   1 to Day 2 of adulthood (unpaired two-tailed Student’s t-test,
   mean ± s.e.m, n = 6 biological replicates per group). d, Seahorse-based
   oxygen consumption of L4 stage N2 worms from the Fig. [287]3f. e,
   Additional integrity scoring parameters of GFP-labeled muscle fibers in
   the worms from the Fig. [288]3g (One-way ANOVA, mean ± s.e.m, n = 6
   animals, 9–31 sarcomeres per group). f, Lifespan of N2 worms treated
   with vehicle or trigonelline starting from Day 4 of adulthood (log-rank
   test, n = 90 animals per group). g, Percentage of paralyzed aged N2
   worms (D11=Day 11) after vehicle or trigonelline treatment started at
   Day 4 of adulthood (n = 3 independent experiments, unpaired two-tailed
   Student’s t-test, mean ± s.e.m). h, Lifespan in N2 worms treated with
   or without NA or trigonelline and fed with control (empty vector; e.v.)
   or nrpt-1 RNAi constructs from Day 1 of adulthood (comparing NA-treated
   worms to e.v., log-rank test, n = 100 animals per group). i, Lifespan
   in N2 worms treated with or without NA or trigonelline and fed with
   control (e.v.) or sir-2.1 RNAi from Day 1 of adulthood (comparing
   NA-treated worms to e.v., log-rank test, n = 100 animals per group). j,
   sir-2.1 mRNA levels by qPCR in D1 adult N2 worms fed with control
   (e.v.) or sir-2.1 RNAi from embryo stage (unpaired two-tailed Student’s
   t-test, mean ± s.e.m, n = 8 biological replicates per group). k,
   sirtuin mRNA levels in N2 worms from publicly available RNA seq
   data^[289]44 (n = 9 replicates per group). l, NAD^+ levels (% control)
   in D1 adult N2 worms treated with trigonelline from embryo stage and
   fed with control (e.v.) or sir-2.1 RNAi constructs (One-way ANOVA,
   mean ± s.e.m, n = 6 biological replicates per group). P: <0.05 (*);
   < 0.01 (**); < 0.001 (***); < 0.0001 (****); n.s., non-significant. For
   all the individual p values, see the Extended Data Fig. 3 Source file.

   [290]Source data

   To translate the functional impact of trigonelline on muscle health
   from nematodes to mammals, we finally supplemented aged mice with
   dietary trigonelline. A 5-day treatment increased the expression and
   activity of mitochondrial oxidative phosphorylation complexes I and II
   in skeletal muscle (Fig. [291]4a–d and Extended Data Fig. [292]4c),
   without influencing mitochondrial abundance measured with
   mitochondrial/nuclear DNA and citrate synthase (Extended Data Fig.
   [293]4a,b). A 12-week dietary trigonelline supplementation increased
   plasma, liver and muscle levels of trigonelline in aged mice (Fig.
   [294]4e,f), with no signs of toxicity (Extended Data Fig. [295]4d,e).
   Body composition was not impacted by the treatment, with no changes in
   lean and fat mass distribution in trigonelline-treated mice (Fig.
   [296]4g and Extended Data Fig. [297]4e), no change of liver and muscle
   mass (Fig. [298]4h and Extended Data Fig. [299]4h,i) and no effects on
   tibialis anterior muscle histology assessed using myofibre size,
   capillary area, glycogen accumulation or fibrosis in tibialis anterior
   and diaphragm muscle (Extended Data Fig. [300]4j–n). We next measured
   NAD^+ in different tissues, where chronic trigonelline exposure
   significantly increased NAD^+ in the liver and kidney (Extended Data
   Fig. [301]4o). Elevation of muscle NAD^+ (Fig. [302]2b and Extended
   Data Figs. [303]2b and [304]4o) and oxidative phosphorylation proteins
   (Fig. [305]4b,d and Extended Data Fig. [306]4p) were flattened in
   chronic versus acute exposure, probably because of long-term
   physiological compensation as observed in clinical studies in muscle
   with dietary administration of other NAD^+ precursors^[307]46,[308]47.

Fig. 4. Trigonelline supplementation enhances mitochondrial activity and
muscle function in aged mice.

   [309]Fig. 4
   [310]Open in a new tab

   a, Mitochondrial complex I activity normalized to citrate synthase
   activity (UCI × UCS^−1) in gastrocnemius muscle of aged mice (20
   months) after a 5-day dietary supplementation of trigonelline
   (unpaired, two-tailed Student’s t-test, n = 7–8 biological replicates
   per group). b, Mitochondrial complex I (NDUFB8) protein levels in the
   same samples as in a (unpaired, two-tailed Student’s t-test, n = 6
   biological replicates per group). c, Succinate dehydrogenase (SDH)
   activity in the quadriceps muscle of the same groups as in a (unpaired,
   two-tailed Student’s t-test, n = 7 biological replicates per group). d,
   Mitochondrial complex II (succinate dehydrogenase [ubiquinone]
   iron-sulphur subunit, mitochondrial (SDHE)) protein levels in the same
   samples as in a (unpaired, two-tailed Student’s t-test, n = 6
   biological replicates per group). e, Plasma trigonelline levels
   measured in aged mice (22–24 months) after 12 weeks of trigonelline
   supplementation (unpaired, two-tailed Student’s t-test, n = 13 and 15
   biological replicates per group). f, LC–HRMS measurement of
   trigonelline levels in gastrocnemius muscle and liver of the same mice
   groups as in e (unpaired, two-tailed Student’s t-test; gastrocnemius:
   n = 5 and 6, liver: n = 13 and 16 biological replicates per group);
   <LOQ, below the level of quantification. g, Lean mass normalized to
   body weight of aged mice after 12 weeks of treatment as in e (unpaired,
   two-tailed Student’s t-test, n = 13 and 15 biological replicates per
   group). h, Tibialis anterior muscle mass of aged mice after 12 weeks of
   treatment as in e (unpaired, two-tailed Student’s t-test, n = 13 and 15
   biological replicates per group). i, Grip strength of aged mice after
   12 weeks of treatment as in e (unpaired, two-tailed Student’s t-test,
   n = 13 and 15 biological replicates per group). j, In situ muscle
   contractility normalized to initial force after supramaximal
   stimulation of the tibialis anterior muscle in young controls and aged
   mice after 12 weeks of treatment as in e (two-way ANOVA followed by
   uncorrected Fisher’s least significant difference tests; n = 11–14
   biological replicates per group). All data are expressed as the
   mean ± s.e.m. with *P < 0.05, **P < 0.01, ***P < 0.001, ****P < 0.0001.
   Individual P values are reported in Fig. 4 of the [311]Source data.

   [312]Source data

Extended Data Fig. 4. Effects of chronic trigonelline supplementation on
physiological and muscle molecular readouts during aging.

   [313]Extended Data Fig. 4
   [314]Open in a new tab

   a, mt/nDNA of gastrocnemius muscle of aged C57BL/6J mice (20 months)
   following a 5 day diet supplementation of trigonelline (unpaired
   two-tailed Student’s t-test, mean ± s.e.m, n = 8 animals per group),
   related to Fig. [315]4a–d. b, Citrate synthase activity and additional
   enzymatic activity of Oxphos subunits from gastrocnemius muscles of the
   same groups as in (a) (unpaired two-tailed Student’s t-test,
   mean ± s.e.m, n = 8 animals per group, 3 samples in control group for
   complex V analyses could not be measured). c, Western blot of SDHB and
   NDFUB8 Oxphos subunits, and Licor staining for total proteins,
   quantified in Fig. [316]4b, d. d,e Plasma aspartate aminotransferase
   (AST) and creatine kinase levels measured in aged C57BL/6J mice (22–24
   months) after a 12 week trigonelline supplementation (unpaired
   two-tailed Student’s t-test, mean ± s.e.m, n = 13 and 15 animals per
   group), related to Fig. [317]4e–j. f, Fat mass composition at the end
   of the intervention study normalized to body weight in the same groups
   as in (d) (unpaired two-tailed Student’s t-test, mean ± s.e.m, n = 13
   and 15 animals per group). g, Liver mass at the end of the study
   normalized to body weight (unpaired two-tailed Student’s t-test,
   mean ± s.e.m, n = 13 and 15 animals per group). h-i, Muscle mass of
   additional muscles at the end of the study normalized to body weight
   (unpaired two-tailed Student’s t-test, mean ± s.e.m, n = 13 and 15
   animals per group). j, Quantification of vascularization via CD31
   immunostaining in TA muscle of the control and trigonelline-treated old
   mice (One-way ANOVA; mean ± s.e.m, n = 12 and 14 animals per group). k,
   Fiber size distribution in TA muscle of the same groups, with an
   average of 1543 ± 13 µm^2 for trigonelline treated group and
   1575 ± 13 µm^2 for control group (Kolmogorov-Smirnov test, n = 13 and
   16 animals per group). l, Average fiber area of TA muscle quantified in
   the same groups (unpaired two-tailed Student’s t-test, mean ± s.e.m,
   n = 13 and 16 animals per group). m, Quantification of PAS glycogen
   staining of TA muscles of the same groups (unpaired two-tailed
   Student’s t-test, mean ± s.e.m, n = 12 and 15 animals per group). n,
   Quantification of Van Gieson (VG) fibrosis staining of TA and diaphgram
   muscles of the same groups (unpaired two-tailed Student’s t-test,
   mean ± s.e.m, n = 12 and 15 animals per group). o, NAD^+ levels
   relative to untreated old mice measured in liver, gastrocnemius and
   kidney tissues after chronic trigonelline administration (unpaired
   two-tailed Student’s t-test, mean ± s.e.m, n = 13 and 15 biological
   replicates per group for liver and muscle, n = 9 and 10 biological
   replicates per group for kidney). p, Western blot and related
   quantification of Oxphos protein levels in gastrocnemius muscle of the
   aged control and trigonelline-treated mice (unpaired two-tailed
   Student’s t-test, mean ± s.e.m, n = 5 biological replicates per group).
   q, Spontaneous fine activity of the aged control and
   trigonelline-treated mice, and of the young reference group (One-way
   ANOVA; mean ± s.e.m, n = 13 aged animals, and 16 animals for young and
   trigonelline groups). r, Peak force normalized to body weight of the
   same groups (unpaired two-tailed Student’s t-test, mean ± s.e.m, n = 13
   and 15 biological replicates per group). P: <0.05 (*); < 0.01 (**);
   < 0.001 (***); < 0.0001 (****); n.s., non-significant. For all the
   individual p values, see the Extended Data Fig. 4 Source file.

   [318]Source data

   The effects of trigonelline on in vivo muscle function were assessed by
   recording spontaneous activity and measuring by grip strength and in
   situ tibialis anterior muscle contraction via electrical stimulation of
   the sciatic nerve to overcome any effects on volition. Chronic
   supplementation of trigonelline significantly increased the grip
   strength of the forelimb muscles in aged mice (Fig. [319]4i) and
   normalized age-related decline of fine spontaneous activity that
   mobilizes the hindlimb and forelimb muscles (Extended Data Fig.
   [320]4q). The maximum tetanic force of the tibialis anterior muscles
   was not impacted in aged trigonelline-supplemented mice (Extended Data
   Fig. [321]4r), probably because tetanic force relies on muscle mass and
   neuromuscular coupling, both of which were not affected by
   trigonelline. However, when the tibialis anterior muscles were
   repeatedly stimulated in situ to assess muscle fatigue from
   high-intensity contractions, the age-related decline of the force was
   attenuated in aged mice treated with trigonelline (Fig. [322]4j). In
   this setting, which mimics a physiological situation where the
   performance of healthy young mice decreases by 10% during the fatigue
   protocol, muscle fatiguability was tripled in aged mice but
   trigonelline prevented approximately 50% of this age-related decline.
   Together, these functional data indicate that chronic trigonelline
   administration mitigates muscle decline during ageing by stimulating
   mitochondria and increasing muscle performance and resistance to
   fatigue during high-intensity contraction.

   Although there is a common trajectory of age-related muscle decline in
   all individuals, the rate and extent of loss of muscle mass and
   strength varies between older individuals; furthermore, the prominent
   factors that drive the inter-individual susceptibility to sarcopenia
   and disability versus healthy ageing are poorly understood for
   therapeutic intervention. Our previous profiling of muscle biopsies
   from patients with sarcopenia versus age-matched healthy controls
   revealed that mitochondrial dysfunction and reduced NAD^+, two
   well-established molecular hallmarks of organismal ageing^[323]11, are
   prominent features of pathological progression and sarcopenia^[324]3.
   In this study, we expanded our initial analysis to a targeted serum
   metabolomic profiling of sarcopenia and discovered that the NA-related
   alkaloid trigonelline is a circulating metabolite correlating with
   muscle strength and gait speed in humans, and declines during
   sarcopenia. Trigonelline is a well-described plant metabolite. Its
   synthesis from NA via methylation confers protective and adaptive
   functions^[325]4,[326]48, without an established reutilization as a
   Preiss–Handler substrate^[327]49,[328]50. Consequently, trigonelline is
   particularly abundant in plant-derived food products, such as coffee
   beans and fenugreek seeds, which have been reported to modulate
   endogenous trigonelline levels in humans^[329]4,[330]51,[331]52.
   Although high coffee consumption has been associated with a lower
   prevalence of sarcopenia in some populations^[332]53, our correlation
   analyses did not detect an association between circulating trigonelline
   and dietary caffeine intake levels in a population from the Middle
   East, possibly because coffee consumption is low in this population.
   Serum trigonelline levels were also not correlated to dietary vitamin
   B[3] intake, suggesting that methylation of dietary vitamin B[3] does
   not directly control endogenous trigonelline. It is also unlikely that
   intake of coffee and niacin directly drives the link between
   trigonelline and sarcopenia as the association of serum trigonelline
   with muscle function was not markedly influenced by correction of
   dietary caffeine and vitamin B[3] intake. Interestingly, food may still
   contribute to trigonelline metabolism as the dietary intake of folate
   and fibre was linked to circulating trigonelline levels. In addition,
   trigonelline can be produced by the microbiome. Trigonelline was
   proposed as a biomarker of metabolic health or physical fitness because
   urinary trigonelline levels decrease during obesity and increase in
   professional athletes^[333]29,[334]30. Therefore, the association of
   trigonelline and sarcopenia probably has multifactorial contributions
   via a complex cross-talk between different food groups and endogenous
   metabolism.

   Our work also describes trigonelline as an NAD^+ precursor and
   demonstrates the benefits of its therapeutic dietary supplementation
   for mitochondrial function, muscle health and mobility during ageing.
   NAD^+ boosting by trigonelline is conserved across different model
   organisms and primary human muscle cells from healthy individuals and
   donors with sarcopenia. Trigonelline treatment in aged mice increased
   different domains of muscle performance that mobilize both leg and arm
   muscles by improving both muscle fatigue and grip strength, which are
   important for healthy physical ageing. While the preclinical tests of
   muscle function used do not always directly translate to human
   performance, fatigue and grip strength are also well-established
   measures of sarcopenia and physical frailty^[335]54,[336]55 and are
   commonly used to assess the pathological impairment of quality of life
   in older people in clinical practice. However, sarcopenia is a
   multifactorial disease where both muscle mass and function decline
   beyond pathological thresholds^[337]7,[338]8. As trigonelline improved
   grip strength and fatigue independently of changes in muscle mass and
   maximal tetanic strength, we conclude that trigonelline cannot reverse
   all causes of sarcopenia and will have to be combined with other
   nutrients that support muscle mass, such as protein, vitamin D or omega
   3 fatty acids, or for the nutritional management of sarcopenia^[339]7.
   At the cellular level, the benefits of trigonelline on muscle
   performance did not arise from structural adaptations in the skeletal
   muscle architecture, such as myofibre cross-sectional area,
   vascularization and fibrosis, but were linked to functional
   improvements of the mitochondrial respiratory activity of complexes I
   and II. Using inhibitors and genetic loss of function in vitro and in
   vivo, we showed that trigonelline is metabolized via the Preiss–Handler
   pathway and requires NAPRT to boost NAD^+, stimulate mitochondrial
   metabolism and increase mobility and lifespan during ageing. Our work
   with tracers and metabolomics also indicates that trigonelline is
   demethylated before entering the flux of NAD^+ biosynthesis, which is
   in line with early biochemical evidence that an enzymatic activity from
   the liver may demethylate trigonelline in mammals^[340]36. However,
   there is no known demethylase that catalyses trigonelline
   demethylation. Although the correlation we discovered between SHMT2
   expression and serum trigonelline, muscle mass and muscle strength
   suggests that trigonelline could cross-talk with the
   S-adenosyl-L-methionine-dependent methyltransferase activity of SHMT2
   and folate/one-carbon metabolism, future studies are required to
   characterize the mechanisms of trigonelline demethylation and the
   identity of the trigonelline demethylase.

   Our results add trigonelline to the list of established dietary NAD^+
   boosters. These molecules, including NR, NA, NAM and NMN, have shown
   preclinical potential for ameliorating ageing and chronic diseases in
   model organisms and have been investigated in recent human clinical
   trials focused on healthy ageing^[341]46, obesity^[342]56,
   mitochondrial disease^[343]57, neurodegeneration^[344]58 or insulin
   sensitivity^[345]47, with different outcomes. Our work highlights that
   the NAD^+-boosting capabilities of different precursors vary across
   tissues and experimental models according to the relative activity of
   different branches of NAD^+ biosynthesis. Trigonelline contributes to
   the NAD^+ pool via a more indirect route than the ribosylated
   precursors NR and NMN, but our comparative studies demonstrate that
   trigonelline has similar cellular and physiological benefits to NA and
   NR in cells or nematodes, most probably because of its higher stability
   than ribosylated precursors. NA has lipid-lowering properties^[346]33
   and provides benefits in mitochondrial myopathies with impaired NAD^+
   salvage^[347]57; however, it has limited tolerability as it induces
   skin flushing through dermal Langerhans cells because of
   GPR109A-dependent vasodilation^[348]41,[349]59. Our results indicate
   that unlike NA, trigonelline does not activate GPR109A at physiological
   and therapeutic levels, and probably provides a more favourable
   therapeutic profile. While trigonelline primarily engages the
   Preiss–Handler pathway across models, in vivo supplementation also
   elevates NAD^+ metabolites from other biosynthetic routes, supporting
   the inter-organ cross-talk between NAD^+ pathways already observed in
   humans and rodents with NR supplementation^[350]46,[351]60. In
   particular, trigonelline increases NR and NMN in the liver or serum
   both in WT and Naprt KO mice; these secondary NAD^+ fluxes can
   contribute to the NAD^+ pool in vivo in the absence of Naprt. This
   highlights the complexity of systemic NAD^+ metabolism modulation and
   the need to study the interplay between NAD^+ fluxes and to run future
   comparative studies with different precursors in specific models and in
   humans. The heterogeneity of NAD^+ biosynthesis and metabolism also
   suggests that some physiological specificity may exist between
   different precursors across organs, supporting additional translational
   applications of trigonelline. Because of its high bioavailability and
   serum stability, trigonelline can reach the brain and impact cognitive
   performance in mice with Alzheimer’s disease^[352]61, therefore holding
   potential for neurocognitive benefits in addition to muscle health. In
   addition, trigonelline can protect from metabolic dysfunction and
   improve glucose tolerance both in mice and humans^[353]62,[354]63.

   In summary, clinical profiling revealed an association between
   trigonelline and muscle health in humans. Our preclinical experiments
   demonstrated that trigonelline is an NAD^+ precursor that optimizes
   mitochondrial function to improve muscle strength and prevent fatigue
   during ageing. Therefore, trigonelline is a nutritional geroprotector
   with therapeutic potential to manage sarcopenia and other age-related
   pathologies.

Methods

Human studies

   Detailed description of the MEMOSA study is available in the study by
   Migliavacca et al.^[355]3. Twenty Chinese male participants aged 65–79
   years with a diagnosis of sarcopenia and 20 healthy age-matched
   controls were recruited in Singapore under the National Healthcare
   Group Domain-Specific Research Board study (no. 2014/01304); each
   participant gave written informed consent. Muscle biopsy samples were
   fully depleted during the study analyses.

   The cohort of the Bushehr nutritional epidemiology study in older
   people consists of 186 older men aged 60 years and older participating
   in the second stage of the Bushehr elderly health programme. They were
   randomly selected from the population-based prospective cohort study
   conducted in Bushehr, a southern province of Iran^[356]31. Grip
   strength was measured using a digital dynamometer with three repeats of
   each hand, and retaining the highest average handgrip strength for the
   strongest hand. The ALMI was measured using DXA (Discovery Wi), with
   the AMLI calculated for each participant as the sum of the upper and
   lower limb lean mass, expressed in kilograms, divided by the height
   square, expressed in metres. Dietary intake assessment was performed
   using a 24-h dietary recall of all food and beverages consumed,
   performed by expert nutritionists^[357]31. Standard reference tables
   were used to convert household portions to grams and quantified using
   the nutritionist IV package, modified for Iranian foods to obtain daily
   energy, nutrient values and servings of foods consumed. For mixed
   dishes, food groups and nutrients were calculated according to their
   ingredients. The study was approved by the Research Ethics Committee of
   the Endocrinology & Metabolism Research, Tehran University of Medical
   Sciences, under reference TUMS.EMRI.REC.1394.0036; and each participant
   gave written informed consent. Local sample analyses were approved by
   the cantonal ethics commission for human research (Commission Cantonale
   d’Éthique de la Recherche sur l’Étre Humain) in Vaud, Switzerland under
   reference 490/14. For both studies, tryptophan and vitaminB[3]
   metabolites were measured in overnight fasting venous blood samples
   using liquid chromatography–tandem mass spectrometry as reported in
   Panahi et al.^[358]64 and Midttun et al.^[359]65.

   Primary myotubes were derived from muscle biopsies from participants in
   the HSSe with approval from the Hertfordshire Research Ethics
   Committee^[360]66. Each participant gave written informed consent.

Human RNA-seq and pathway analysis

   RNA-seq of vastus lateralis muscle biopsies of the MEMOSA Singapore
   Sarcopenia Study (SSS) study was performed on libraries generated from
   250 ng of total RNA using paired-end sequencing on an Illumina HiSeq
   2500 sequencer at a depth more than 75 million reads per sample, as
   described previously^[361]3. All statistical analyses of the
   transcriptomics data were performed using R v.3.3.3 and relevant
   Bioconductor packages (for example, edgeR v.3.16.5). Briefly, after
   excluding one sample with an abnormally low percentage of uniquely
   mapped reads and after removing genes with a mean expression lower than
   20 reads, data were normalized by the trimmed mean of M-values as
   implemented in the edgeR^[362]67 function calcNormFactors. Normalized
   skeletal muscle mRNA expression was associated with the serum level of
   trigonelline using Spearman rank correlation. The full dataset of genes
   expressed in skeletal muscle was used for pathway enrichment. Pathway
   enrichment analysis was performed using the Molecular Signature
   Database v.5.2 collections H (hallmark gene sets) and the mean-rank
   gene set enrichment test^[363]68 to assess whether an annotated set of
   genes was enriched in genes associated with serum levels of
   trigonelline. For Extended Data Table [364]5, the list of genes with
   mRNA expression correlating with serum trigonelline was filtered for
   genes annotated with demethylase or methyltransferase activity.

Cell cultures and reagents

   Human primary myoblasts from male donors (catalogue no. CC-2580, Lonza)
   were seeded in 96-well plates at a density of 12,000 cells per well in
   skeletal muscle growth medium (SKM-M0, AMSBIO). After 1 day,
   differentiation was performed by changing to DMEM/F12 (Thermo Fisher
   Scientific) supplemented with 2% horse serum for 4 days. Primary human
   myoblasts from male donors from the HSSe cohort (n = 3 with sarcopenia
   and n = 3 controls) were seeded in 96-well plates coated with Matrigel
   at a density of 15,000 cells per well in myoblast proliferation medium
   (DMEM, 20% FCS, 10% horse serum, 1% penicillin-streptomycin, 1% chick
   embryo extract). After 48 h, differentiation was induced by changing to
   differentiation medium (DMEM, 2% horse serum, 1%
   penicillin-streptavidin). HepG2 cells (catalogue no. HB-8065, ATCC),
   were maintained in exponential growth phase in DMEM supplemented with
   10% FCS, at 37 °C in a humidified atmosphere of 5% CO[2]. C2C12
   myoblasts (catalogue no. CRL-1772, ATCC) were cultured in a 5% CO[2]
   incubator in high-glucose DMEM (Gibco) containing 10% FCS, and were
   differentiated into myotubes in high-glucose DMEM containing 2% horse
   serum when cell density reached 80–90% at 37 °C. Primary myotubes from
   aged male C57BL/6JRj mice (24 months; Janvier Labs) were generated from
   freshly sorted muscle stem cells isolated using flow cytometry as
   described previously^[365]69. Briefly, hindlimb muscles were rapidly
   collected, minced and digested with 2.5 U ml^−1 Dispase II
   (Sigma-Aldrich), 0.2% Collagenase B (Sigma-Aldrich) and 5 mM MgCl[2] at
   37 °C. The preparation was then filtered sequentially through 100-μm
   and 30-μm filters and cells were incubated at 4 °C for 30 min with
   antibodies against CD45 (1:25 dilution, catalogue no. MCD4528,
   Invitrogen), CD31 (1:25 dilution, catalogue no. RM5228, Invitrogen),
   CD11b (1:25 dilution, catalogue no. RM2828, Invitrogen), CD34 (1:60
   dilution, catalogue no. 560238, BD Biosciences), Ly-6A/E (1:150
   dilution, catalogue no. 561021, BD Biosciences) and α7-integrin (1:30
   dilution, catalogue no. FAB3518N, R&D Systems). Muscle stem cells
   identified as CD31^−CD11b^−CD45^−Sca1^−CD34^+integrin α7^+ were
   isolated with a Beckman Coulter Astrios Cell sorter.
   Fluorescence-activated cell sorting-isolated muscle stem cells were
   plated onto gelatin-coated 384-well plates at a density of 600 cells
   per well in DMEM supplemented with 20% heat-inactivated FCS (catalogue
   no. 16140063, Thermo Fisher Scientific), 10% horse serum (catalogue no.
   26050088, Thermo Fisher Scientific), 2.5 ng ml^−1 basic fibroblast
   growth factor (catalogue no. PMG0035, Thermo Fisher Scientific), 1%
   penicillin-streptomycin (catalogue no. 15070063, Thermo Fisher
   Scientific) and 1% l-glutamine (catalogue no. 25030149, Thermo Fisher
   Scientific). Medium was refreshed the next day and cells were grown for
   4 days until confluency. Cells were then differentiated for 2 days to
   myotubes in differentiation medium (DMEM, catalogue no. 11995065,
   Thermo Fisher Scientific) supplemented with 2% horse serum and 1%
   penicillin-streptomycin. Treatments were performed on day 4 of
   differentiation for 24 h. IM-PTEC cells isolated from immorto
   mice^[366]70 were a gift from A. Tammaro (Amsterdam UMC). Cells were
   maintained in DMEM/F12 medium (Gibco) with 10% FCS (Gibco), 5 μg ml^−1
   insulin and transferrin, 5 ng ml^−1 sodium selenite (Gibco),
   40 pg ml^−1 triiodothyrionine (Sigma-Aldrich), 36 ng ml^−1
   hydrocortisone (Sigma-Aldrich) and 20 ng ml^−1 epidermal growth factor
   (Sigma-Aldrich) with l-glutamine and antibiotics (both from Gibco) and
   mouse interferon-y (IFN-y) (10 ng ml^−1; ProSpec) at 33 °C in 5% (v/v)
   CO[2]. Before being seeded for the experiments, cells were grown under
   restrictive conditions (37 °C 5% (v/v) CO[2], in the absence of IFN-γ)
   for 7 days to downregulate SV40 large T antigen activity. Cell lines
   were regularly tested using a Rapid Mycoplasma Detection Kit
   (MycoGenie, Assay Genie).

   Cells were treated with trigonelline iodide (catalogue no.
   MolPort-003-944-936, Molport) after internal quality control using mass
   spectrometry, showing more than 99.7% purity, NR chloride (ChromaDex),
   NA (catalogue no. N0761, Sigma-Aldrich), NAM (catalogue no. 72340,
   Sigma-Aldrich), NMN (catalogue no. N3501, Sigma-Aldrich), 2-OHNA
   (catalogue no. 251054, Sigma-Aldrich) and FK866 (catalogue no. F8557,
   Sigma-Aldrich) for the desired time. Unless otherwise stated,
   trigonelline treatment was performed at 1 mM. 2-OHNA and FK866 were
   used at 1 mM and 100 nM, respectively in all experiments, unless
   indicated otherwise. Hoechst [367]H33342 solution (Invitrogen, Thermo
   Fisher Scientific) and JC-10 (catalogue no. ENZ-52305, Enzo Life
   Sciences) were used to assess mitochondrial function. Isotopically
   labelled trigonelline iodide ([^13C,^2H[3]]-trigonelline M + 4) was
   labelled with one ^13C on the carbonyl group and three ^2H on the
   nitrogen of the pyridine ring through custom synthesis and used in
   vitro and in vivo at the mentioned concentrations. Briefly,
   [carbonyl-^13C]-NA (0.1 g, 0.8 mmol) was stirred in anhydrous ethanol
   (5 ml), followed by adding 200 μl (0.46 g, 3 mmol) of ^2H[3]-methyl
   iodide at 23 °C. The mixture was stirred under reflux at 45–48 °C in an
   oil bath for 2 days. The oil bath was removed and the reaction mixture
   was allowed to cool down to room temperature. Ethanol was evaporated on
   a rotary evaporator. Yellow powder residue was first washed with
   ethanol (2× 1 ml) and then with diethyl ether (2 ml). The product was
   additionally dried under vacuum at 23 °C to give yellow solid (0.159 g,
   74%; MS: 142.14 [M-I]+).

NAD^+ measurement

   Cellular NAD^+ levels were assessed using the BioVision NAD^+/NADH
   Quantitation Colorimetric Kit (catalogue no. k337-100). Total NAD^+ was
   normalized to the total number of cells. Quantification of NAD^+ in
   tissue samples was performed using an enzymatic method adapted from
   Dall et al.^[368]71 with normalization to tissue weight. Analysis of
   the NAD^+ metabolome in cells or in vivo samples was performed using
   high-resolution liquid chromatography–mass spectrometry as described
   below.

Cellular assays

Cell death assays

   Kinetic experiments of apoptosis were performed with the Incucyte ZOOM
   instrument (Essen BioScience). Cells were incubated with Incucyte
   Annexin-V Green (catalogue no. 4642), according to the supplier’s
   instructions. Four images per well were collected at the indicated time
   using a 10× objective and bandwidth filters (excitation: 440/80 nm;
   emission: 504/44 nm).

Mitochondrial function

   For mitochondrial potential assessment, cells were seeded into a
   96-well Sensoplate (catalogue no. 655090, Greiner) at a density of
   12,000 cells per well in SKM-M. After 1 day, differentiation was
   induced by a medium change for 4 days; mitochondrial potential was
   performed at the desired time and with the desired treatment. Briefly,
   cells were incubated for 30 min in Krebs buffer with Hoechst staining
   and JC-10. Images were acquired using ImageXpress (Molecular Devices)
   using a 10× objective. The following filters were used: JC-10, FITC
   Filter Cube/TRITC Filter Cube; Hoechst, 4,6-diamidino-2-phenylindoleI
   Filter Cube. The total intensity of both FITC and TRITC fluorescence
   was recorded for each cell and was used to calculate a cellular
   fluorescence ratio: ratio per cell = log[2] (∑ pixel intensity TRITC/∑
   pixel intensity FITC). Once segmentation was completed, results were
   analysed with the KNIME software v.4.3.1.

   The bioenergetic profiles of the cells were analysed using an XF96
   extracellular flux analyzer (Seahorse Bioscience) as described.
   Briefly, cells were seeded in XF96 Cell Culture Microplates and the OCR
   was measured in Krebs buffer (NaCl 135 mM, KCl 3.6 mM, NaH[2]PO[4]
   0.5 mM, MgSO[4] 0.5 mM, HEPES 10 mM, NaHCO[3] 5 mM) supplemented with
   10 mM glucose, 10 mM pyruvate and 2 mM glutamine. For substrate-driven
   OCR measurement in permeabilized HSMMs, the assay was performed in
   Seahorse XF Plasma Membrane Permeabilizer (Agilent Technologies)
   according to the manufacturer’s instructions. Briefly, cells were
   seeded in XF96 Cell Culture Microplates and the OCR was measured in
   mitochondrial assay solution (MAS buffer; mannitol 220 mM, sucrose
   70 mM, KH[2]PO[4] 10 mM, MgCl[2] 5 mM, HEPES 2 mM, EGTA 1 mM, BSA 0.2%
   (w/v)) supplemented with ADP.

NAPRT knockdown in cells

   To knockdown NAPRT in myotubes, 8,000 cells per well were seeded
   overnight in SKM-M medium. Cells were then infected with NAPRT shRNA or
   scrambled shRNA adenoviruses (SIRION Biotech) at 200 multiplicity of
   infection; cells were incubated for 48 h before initiating
   differentiation of myoblasts into myotubes. To check knockdown
   efficiency, cells were lysed in TriPure RNA reagent. Total RNA was
   transcribed to complementary DNA (cDNA) using the QuantiTect Reverse
   Transcription Kit (QIAGEN); mRNA expression was analysed using the
   LightCycler 480 instrument (Roche Diagnostics) and LightCycler 480 SYBR
   Green I Master Reagent (Roche Life Science). See Extended Data Table
   [369]6 for the list of primers used.

Extended Data Table 6.

   qPCR primers

   [370]graphic file with name 42255_2024_997_Tab6_ESM.jpg
   [371]Open in a new tab

   List of qPCR primers used to detect C. elegans, mice and human gene
   expression in this study (5′ > 3′).

GPCR agonist assay

   The GPR109A agonist assay was performed using the PathHunter β‐arrestin
   enzyme fragment complementation technology (Eurofins). Compounds were
   tested in duplicate in agonist mode and data were normalized to the
   maximal and minimal response observed in the presence of positive
   control and vehicle, respectively.

NAD^+ precursor stability

   The stability of the different precursors was assessed in human serum
   (catalogue no. H4522, Sigma-Aldrich) at 37 °C at the indicated time
   points. A liquid–liquid extraction was adopted from Giner et
   al.^[372]72 to assess the level of precursors or intermediates using
   the analytical methods described below.

Animal studies

   Studies and procedures in WT mice were approved by the Nestlé Ethical
   Committee (ASP-19-03-EXT), the Office Vétérinaire Cantonal Vaudois
   (VD2770 and VD3484) and the Animal Ethics Committee at The University
   of Melbourne (1914961.2). Naprt KO animal studies were approved by the
   Animal Experiment Committee at the University of Toyama (approval no.
   A2022MED-19) and were performed in accordance with the Guidelines for
   the Care and Use of Laboratory Animals at the University of Toyama,
   which are based on international policies.

   In vivo treatments were performed with trigonelline monohydrate (Laurus
   Labs) after internal quality control using MS, showing more than 99.9%
   purity, or [^13C,^2H[3]]-trigonelline M + 4 iodide synthesized by the
   authors M.V.M. and M.E.M. for this study. Trigonelline was administered
   orally at a dose equimolar to 300 mg kg^−1 of anhydrous trigonelline.
   For the in vivo tracer experiments, 12-week-old C57BL/6JRj male mice
   were given an acute dose of 300 mg kg^−1 labelled
   [^13C,^2H[3]]-trigonelline M + 4 iodide by oral gavage; after 2 h and
   24 h, blood, urine, muscle and liver were sampled. For chronic
   trigonelline supplementation, mice were fed with a standard chow diet
   alone or supplemented before pelleting with trigonelline monohydrate
   (Laurus Labs), at a dose delivering an average exposure of 300 mg kg^−1
   per day. Twenty-month-old aged C57BL/6JRj male mice were fed control or
   trigonelline diet (catalogue no. s8189, Ssniff) for 5 days, while young
   12-week-old control and 20-month-old aged C57BL/6J male mice (The
   Jackson Laboratory) were fed control or trigonelline diet (AIN93M,
   Specialty Feeds) for 12 weeks. In the 12-week study, phenotypic
   characterization included body composition, using a whole-body
   composition analyser (LF50, Bruker), spontaneous physical activity of
   fine and ambulatory movements measured in Promethion metabolic cages
   (Sable Systems International), and grip strength assessed 1 week before
   the end of the study using a grip strength metre recording maximal
   force at peak tension before releasing the grasp. At the end of the
   study, tibialis anterior contractile muscle function was assessed under
   anaesthesia as described previously^[373]73,[374]74. Briefly, maximum
   isometric tetanic force was determined from the plateau of a complete
   frequency–force relationship. Assessment of contraction-induced fatigue
   was determined by maximally stimulating muscles for an isometric
   contraction once every 4 s for 4 min and normalizing the decline of
   force to the baseline value. Force recovery was assessed after 5 min
   and 10 min rest after the initial stimulation period. A blood sample
   was subsequently obtained via cardiac puncture; after cardiac excision,
   skeletal muscles (tibialis anterior, extensor digitorum longus, soleus,
   plantaris, gastrocnemius, quadriceps, diaphragm strips) and organs
   (liver, kidneys) were surgically excised, weighed and snap-frozen for
   biochemical analysis or embedded and frozen in isopentane cooled in
   liquid nitrogen for immunohistochemical analysis. Plasma parameters
   were measured using the VetScan Equine Profile Plus.

   For the Naprt KO study, animals were kept under a controlled
   temperature and humidity (25 °C, 50%) with standard light condition (a
   12:12 h light–dark cycle) with free access to water and standard chow
   diet (CLEA Japan). Naprt KO mice of mixed sexes were obtained by
   crossing the heterogenic C57BL/6N Naprt KO mice described
   previously^[375]40 and treated with custom-synthesized trigonelline
   iodide at 8 weeks of age. Two hours later, tissues were collected and
   immediately frozen in liquid nitrogen and kept in −80 °C.

Histological analyses

   After cryosectioning, muscle sections were stained with haematoxylin &
   eosin for assessment of muscle architecture, CD31 to visualize
   capillaries, Van Gieson’s stain to identify collagen and fibrosis, SDH
   activity as a general marker of mitochondrial (oxidative) activity, and
   periodic acid–Schiff stain for glycogen content^[376]75. The muscle
   fibre cross-sectional area was measured after immunolabelling of
   laminin; fibre type was identified by staining myosin heavy chain I and
   myosin heavy chain IIa. Digital images of stained sections (four images
   per muscle section) were obtained using an upright microscope (20×
   objective) with a camera (Axio Imager D1, ZEISS) and AxioVision AC
   software (AxioVision AC, release 4.7.1, ZEISS) for acquisition.
   Myofibre cross-sectional area was quantified as described
   previously^[377]76,[378]77.

RNA and immunoblot analyses in rodent tissues

   For qPCR of tissues from the Naprt KO study, RNA extraction was
   performed using the TRI Reagent (Sigma-Aldrich). The ReverTra Ace qPCR
   RT Master Mix with genomic DNA Remover (TOYOBO) was used to synthesize
   cDNA. Real-time PCR was performed using the THUNDERBIRD SYBR qPCR Mix
   (TOYOBO) on the Thermal Cycler Dice Real Time System II (Takara Bio).
   mRNA was quantified using the ∆∆^Ct method against Rpl13a as the
   reference gene. See Extended Data Table [379]6 for the list of primers
   used.

   Oxidative phosphorylation complex expression was measured in isolated
   mitochondria by immunoblot using an antibody cocktail (1:250 dilution,
   catalogue no. 45-8099, Invitrogen) and anti-α-tubulin antibody (1:1,000
   dilution, catalogue no. T6074, Sigma-Aldrich). Equal protein load
   (20 µg per well) and consistent gel transfer was verified by Revert
   Total Protein Stain (LI-COR). Immunoblots were imaged using a LI-COR IR
   imager and quantified with Image Studio Lite (v.5.0, LI-COR).

Metabolomics analyses in cells and rodent tissues

   Metabolomics analyses were conducted using a liquid–liquid extraction
   method adapted from Giner et al.^[380]72. For cell extraction, cells
   were scraped and extracted in a cold mixture of
   methanol:water:chloroform (5:3:5 (v/v)). Tissue extraction was
   performed with metal beads in pre-cooled racks (−80 °C) using a tissue
   mixer (TissueLyser II, QIAGEN). Whole blood and urine were directly
   extracted in cold methanol:water:chloroform (5:3:5 (v/v)). Isotopically
   labelled internal standards, including fully labelled ^13C yeast
   extract and nicotinamide-d4, were included for data normalization.
   Additionally, isotopically labelled acyl-carnitines (NSK-B, Cambridge
   Isotope Laboratories) were added for normalization in cell
   metabolomics. After centrifugation, samples yielded an upper phase
   containing polar metabolites, a lower phase containing apolar
   metabolites, and a protein layer in between. The upper phase was dried
   and dissolved in 60% (v/v) acetonitrile:water for analysis. Protein
   layers from cells, liver and muscle samples were quantified using a
   bicinchoninic acid assay (Thermo Fisher Scientific) for normalization.
   HRLC–MS spectrometry was performed using hydrophilic interaction liquid
   chromatography (HILIC) analytical columns, such as HILICON iHILIC
   Fusion(P) or ZIC-pHILIC columns^[381]77,[382]78. Separation was
   achieved using a linear solvent gradient with solvent A (H[2]O with
   10 mM ammonium acetate and 0.04% (v/v) ammonium hydroxide, pH
   approximately 9.3) and solvent B (acetonitrile). The eluting NAD^+
   metabolites were analysed using an Orbitrap Fusion Lumos mass
   spectrometer (Thermo Fisher Scientific) with a heated electrospray
   ionization source. Data processing and instrument control were
   performed using the Xcalibur v.4.1.31.9 software (Thermo Fisher
   Scientific). Quantitative trigonelline measurement in tissues was
   performed using the same liquid–liquid extraction method, while body
   fluids (plasma and urine) were extracted using ACN:MeOH:H[2]O (−20 °C)
   as described by Li et al.^[383]79. A labelled amino acid mix (NSK-A1,
   Cambridge Isotope Laboratories) served as an internal standard.
   Leucine-d4 was used as an internal standard for normalization.
   Trigonelline concentrations were determined using calibration curves
   ranging from 10 µM to 1,000 µM. For the tracer experiments, the same
   methods were used and the relative abundance and enrichment of
   isotopologues of NAD metabolites were calculated by dividing the area
   of each isotopologue by the sum area of all isotopologues. In the Naprt
   KO rodent study, metabolite extraction and NAD^+ metabolomics were
   performed as described previously^[384]80. Tissues were grounded in a
   50% methanol 50% water mixture using a multi-bead shocker (Yasui
   Kikai). Metabolites were analysed using an Agilent 6460 Triple Quad
   Mass Spectrometer coupled with an Agilent 1290 HPLC system.
   Trigonelline and other NAD^+ metabolites were detected using specific
   transitions; data analysis was conducted using the MassHunter
   Workstation Quantitative Analysis software (Agilent Technologies).

C. elegans experiments

   WT hermaphrodite Bristol worms (N2) and GFP-tagged myosin worms^[385]81
   were treated with trigonelline chloride (catalogue no. T5509,
   Sigma-Aldrich) and NR chloride from day 1 of adulthood. Nematodes were
   cultured at 20 °C on nematode growth medium (NGM) agar plates seeded
   with Escherichia coli strain OP50. Trigonelline and the other
   precursors were added to the NGM medium at a final concentration of
   1 mM just before pouring the plates, unless otherwise stated. An
   enzymatic method adapted from Dall et al.^[386]71 was used to measure
   NAD^+ content in worms. Fifty to 100 worms were collected per
   biological replicate and NAD^+ levels were normalized on the protein
   content. The bacterial feeding RNAi experiments were carried out as
   described^[387]82. RNAi was performed using the clones sir-2.1
   (R11A8.4) and nprt-1 (Y54G2A.17) from Geneservice.

Lifespan

   Worm lifespan tests were performed using 90–100 animals per condition
   and scored manually every other day, as described previously^[388]83.
   Treatments and experimental measurements were started on day 1 of WT N2
   worm adulthood, in a regimen of chronic exposure during the entire
   study. Statistical significance was calculated using the log-rank
   (Mantel–Cox) method.

Mobility assay

   C. elegans spontaneous mobility was measured using the Movement Tracker
   software (v.1)^[389]84. For paralysis scoring, 45–60 worms per
   condition were manually scored for mobility after poking. Worms that
   did not respond to any repeated stimulation were scored as dead.
   Results are representative of data obtained from at least two
   independent experiments.

RNA analyses

   A total of approximately 6,000 worms per condition, divided into six
   biological replicates, was recovered in M9 buffer from the NGM plates
   at day 2 of adulthood and lysed in the TriPure RNA reagent. Total RNA
   was transcribed to cDNA using the QuantiTect Reverse Transcription Kit.
   Expression of selected genes was analysed using the LightCycler 480
   system and LightCycler 480 SYBR Green I Master Reagent. For worms, two
   housekeeping genes were used to normalize the expression data, that is,
   actin (act-1) and peroxisomal membrane protein 3 (pmp-3). See Extended
   Data Table [390]6 for the list of primers used.

Muscle integrity imaging

   Imaging of muscle structure was performed on RW1596 myo-3 (st386)
   worms^[391]55. Trigonelline treatment started on day 1 of adulthood, in
   a regimen of chronic exposure until day 11. Worms were immobilized with
   a 7.5-mM solution of tetramisole hydrochloride (Sigma-Aldrich) in M9
   and mounted on 2% agarose pads on glass slides. Confocal images were
   acquired with Leica SP8 inverse STED 3X (Leica Microsystems) under
   non-saturating exposure. Myofibre integrity scoring was quantified as
   described by Dhondt et al.^[392]85 on a total of six worms per group
   and 9–31 muscle cells per group.

Oxygen consumption assays

   Oxygen consumption was measured in N2 worms treated from embryo to L4
   with trigonelline using the Seahorse XF96 system (Seahorse Bioscience)
   as described previously^[393]86. Respiration rates were normalized to
   the number of worms in each well and averaged over five measurements.

Mitochondrial DNA quantification

   Worms were lysed in 2 μl lysis buffer (50 mM KCl, 10 mM Tris pH 8.3,
   2.5 mM MgCl[2], 0.45% NP-40, 0.45% Tween 20, 0.01 gelatin, 100 μg ml^−1
   proteinase K (added before use)) and heated at 65 °C for 60 min
   followed by inactivation at 90 °C for 15 min. Worm lysates were diluted
   50× with diethyl pyrocarbonate water for the SYBR Green qPCR reaction
   using a LightCycler 480 SYBR Green I Master Kit assay. Relative values
   for nd-1 and act-3 (Extended Data Table [394]6) were compared in each
   sample to generate a ratio representing the relative level of
   mitochondrial DNA per nuclear genome. Experiments were performed on at
   least ten independent biological samples.

Statistics

   All statistical analyses were performed using Prism v.9 (GraphPad
   Software) or R as described in the figure legends. The statistical
   methods for the transcriptomic analyses of muscle biopsies have been
   reported previously^[395]3. Data distributions plotted as box plots
   represent the 25th percentile, the median and 75th percentile, with the
   whiskers extending from the quartiles to the smallest or biggest value
   within 1.5 times the interquartile range. Associations between two
   continuous variables were determined using Spearman rank correlations.
   For statistical comparisons of two conditions, a two-tailed, unpaired
   Student’s t-test was used. For comparisons of more than two groups,
   data were analysed with a one-way ANOVA followed by Šídák’s multiple
   comparisons post hoc test or a two-way ANOVA, unless specified
   otherwise in the figure legends. The normality of each readout was
   assessed based on historical values of the laboratory for this readout.
   All data represent the mean ± s.e.m. NS indicates results that were not
   statistically significant; *P < 0.05, **P < 0.01, ***P < 0.001 and
   ****P < 0.0001 were considered statistically significant.

Reporting summary

   Further information on research design is available in the [396]Nature
   Portfolio Reporting Summary linked to this article.

Supplementary information

   [397]Reporting Summary^ (4.5MB, pdf)

Source data

   [398]Source Data Fig. 1^ (34.4KB, xlsx)

   Individual values and statistics.
   [399]Source Data Fig. 2^ (44.4KB, xlsx)

   Individual values and statistics.
   [400]Source Data Fig. 3^ (42.3KB, xlsx)

   Individual values and statistics.
   [401]Source Data Fig. 4^ (38.3KB, xlsx)

   Individual values and statistics.
   [402]Source Data Extended Data Fig. 1^ (67.6KB, xlsx)

   Individual values and statistics.
   [403]Source Data Extended Data Fig. 2^ (68.1KB, xlsx)

   Individual values and statistics.
   [404]Source Data Extended Data Fig. 3^ (40.5KB, xlsx)

   Individual values and statistics.
   [405]Source Data Extended Data Fig. 4^ (36.6KB, xlsx)

   Individual values and statistics.
   [406]Source Data Extended Data Fig. 4^ (1.4MB, tif)

   Unprocessed immunoblots.

Acknowledgements