Scalable syntheses of traceable ribosylated NAD⁺ precursors

Introduction

Nicotinamide adenine dinucleotide (NAD⁺, Figure 1) and its phosphorylated and reduced forms, are vitamin B3-derived dinucleotides, redox coenzymes in hydride transfers, reactions for which the nucleosidic nature of the dinucleotide is retained.^1–3 NAD⁺ is also a substrate for a range of non-redox enzymes, which include sirtuins, adenosine diphosphoribosyl transferases (ARTs), pyrophosphorylases and glycohydrolases, reactions in which the dinucleotidic scaffold is lost.⁴ The physiological roles of these classes of NAD⁺ consuming enzymes range from metabolism regulation and glucose use to DNA repair, apoptosis and cell signalling.^{5, 6} During the process of aging and in several disease models, the demand on the NAD⁺ pool far exceeds its availability. This can occur as a consequence of changes in its biosynthesis from nutritional precursors, in the intracellular recycling of its metabolites or in its conversion to biosynthetically non-recyclable metabolites.⁷

Several research groups have shown that in mammals, depletion in NAD⁺ intracellular levels can be restored by the oral administration of a range of NAD⁺ precursors, which include nicotinic acid (NA),^8–10 nicotinamide (Nam)¹¹, nicotinamide riboside (NR),^{12, 13} nicotinic acid riboside (NAR), nicotinamide mononucleotide (NMN)¹³ and more recently dihydronicotinamide riboside (NRH) (Figure 1).¹⁴

While NA and Nam are long-standing vitamin B3 supplements,^{9, 15} it is only recently that the specific pharmacological effects of supplementation with NR or NMN have been positively associated with a wider range of metabolic diseases in animal models, with both NR^16–18 and NMN^{13, 19, 20} shown to either increase NAD⁺ levels via Nicotinamide Riboside Kinase (Scheme 1, NRK) both in cell and mice models^{21, 22}or via a dedicated transporter.²⁰ Dietary supplementation with NR, one of the B3 vitamins detected in mammalian breast milk,²³ has been directly correlated to an increase in circulating NAD⁺ in humans,²⁴ rendering this class of NAD⁺ precursors attractive candidates for applications in health and nutrition.^25–27 Nicotinic acid riboside is also a precursor to NAD⁺²⁸, which employs the NRK pathway to generate nicotinic acid adenine dinucleotide (NAAD), substrate of NAD synthase (NADS) for its conversion of NAD⁺ (Scheme 1). NAR was shown to be particularly valuable in studying neuronal degeneration²⁹. The reduced form of NR (6, NRH) is also a precursor of NAD⁺,¹⁴ and this via its putative intracellular conversion to NADH, the reduced form of NAD⁺. Additionally, NRH is a substrate for the detoxifying enzyme NRH:quinone oxidoreductase 2 (NQO2)³⁰. As such, NRH offers potentially additional physiological functions to that of its oxidised form, NR.

Overall, the effects of these NAD⁺ precursors on intracellular NAD⁺ levels have received much attention, with advances in the field having been facilitated tailor-made isotopologues used in combination with mass spectrometry to measure metabolic fluxes, biodistribution and pharmacokinetic parameters^{21, 22, 31–35}. Surprisingly, the effects of these NAD⁺ precursors on specific organelles remain elusive, and much remains to be investigated to establish how each precursor modulates cofactors’ sub-cellular distribution^{33, 36}.

To unravel the complexity of NAD⁺ biology, each NAD⁺ precursors must be readily accessible. There are numerous reported syntheses of NAD⁺ precursors³⁷ including that of isotopically labelled derivatives⁷. However, they often suffer of average conversion yields and require extensive purification sequences, which promote product’s decomposition. This limits the value of these synthetic approaches when applied to costly isotopically labelled synthetic reagents. This is particularly relevant when generating on large gram scales isotopically labelled NAD⁺ precursors, used in tracers’ experiments for animal studies. Critically, since the synthetic by-products are themselves NAD⁺ precursors, access to materials of the highest purity is a requirement when used in the profiling of the NAD⁺ metabolome and in flux experiments (Scheme 1). Therefore, we thought to develop synthetic strategies which minimize side-product formations, optimize molarity ratios between reagents and maximize conversion. Here, we report such efficient syntheses for each one of the ribosylated species and related isotopologues.

Results

The synthesis of the ribosylated nicotinamide derivatives employed solvent-enabled ball-milling (Scheme 2; BM: ball milling) conditions which are readily scalable. Following silylation (HMDS, (NH₄)₂SO₄ or TMSCl) of Nam and removal of excess silylating reagents under reduced pressure, molar equivalency of silylated nucleobase (2a) was added to a suspension tetraacetate riboside (1) in a 1.5 molar equivalent amount of dichloromethane and subjected to standard Vorbrüggen glycosylation under mechanochemistry conditions. The conversion was stereospecific and complete. After removal of the volatiles, triacetate nicotinamide riboside triflate (3a) was obtained as a solid, with the only admixtures being residual acetic acid and trimethylsilyl acetate.

The subsequent deprotection of the triacetylated nicotinamide riboside was conducted at low temperature (<−20°C) in absence of moisture to minimise cleavage of the glycosidic bond. The triflate salt form of NR triacetate (3a) was placed in a sealable pressure tube and dissolved in anhydrous methanol. The tube, kept under Ar, was cooled to −78°C and dry ammonia gas was bubbled in the solution for 5min. The tube was then sealed and stirred until placed at −20°C, usually for not less than 120 hrs. Although being prolonged, such reaction times resulted in the complete removal of the acetyl groups and production of nicotinamide riboside triflate 5a. The reaction progression may be monitored by ¹H NMR. However, ammonia must be removed under vacuum prior to the sample being dissolved in the deuterated solvent. NR triflate (NR^+-OTf) is a very hydroscopic material which is difficult to isolate in the solid form, but can be converted to NR⁺Cl⁻, either by ion exchange chromatography or by liquid-liquid extraction^{38, 39}.

To generate the reduced form NRH (6a), the triacetylated nicotinamide riboside (3a) was re-suspended in a minimum volume of ethyl acetate or DCM and thoroughly mixed with a saturated and freshly prepared aqueous solution of NaHCO₃ and Na₂S₂O₄. After 15–20min, the organic layer, which contained the reduced triacetate nucleoside (4a) was isolated. Removal of the acetyl groups using methanol and catalytic amount of K₂CO₃ and generation of NRH (6a) was greatly enhanced in terms of rates, yields and purity when conducted under mechanochemical conditions. This sequence enables the preparation of NRH virtually free of residual triflate, carbonate and thionite salts. Of note, access to NRH from commercially available NR salts can be achieved by treatment of the NR precursor with sodium dithionite and potassium carbonate in aqueous solution followed by a purification or by heterogeneous catalysis⁴⁰.

The efficient synthesis of the ribosylated nicotinic acid derivatives (5b) and (6b) was achieved via a similar route. Unlike NR, the triacetate NAR (3b) is a zwitterion and this property was used to generate this molecule free of triflate salts. However, care had to be taken to maintain these derivatives in a zwitterionic form to minimize decomposition. Triacetate NAR could then be readily deprotected under conditions similar to that of NR triflate or reduced under conditions similar to that of triacetate NRH. However, unlike triacetate NRH (4a), triacetate NARH (4b) is very easily oxidized back to (3b), either when in solution or when stored as a solid at low temperature. As such, triacetate NARH should be prepared as required and use rapidly. NARH (6b) is readily generated from (4b) at room temperature in presence of a 1.1 molar amount of 25 wt.% MeONa in methanol but is itself prone to oxidation to NAR (5b).

Unlike canonical nucleosides, vitamin B3 derived nucleosides, nucleotides and dinucleotides are temperature and pH sensitive and at times even air sensitive. While differences between reduced and oxidized forms of the nicotinamide dinucleotides have long been established⁴¹, the synthetic requirements that were implemented for the isolation of NR and NAR with minimal decomposition to their respective pyridinyl parent is indicative that NR and NAR possess somewhat different stability profiles with respect to pH and temperature. Therefore, we sought to establish how each one of these nucleosides responded to a transient pH adjustment of an aqueous solution followed by a long incubation period (Supp. Fig. 1) and we found that NR was more labile to transient basic conditions than NAR and that NAR had a wider pH stability range.

In order to investigate the biosynthetic pathways that these precursors employ when converted intracellularly to NAD⁺ and its related metabolites, we developed a range of isotopologues and isotopomers of Nam, NA, NR, NAR and NRH by applying the chemistry described above. Quantification by liquid-chromatography coupled to mass spectrometry (LC-MS) of the NAD⁺ metabolome including that incorporating isotopically labelled units reflects the incorporation and biodistribution of isotopically labelled species. ²H or ¹³C labels in the riboside unit can be used to monitor the conversion and turn-over of the ribosylated species in cells, in organelles, in tissues or in organs^{7, 20, 21, 33}. Furthermore, suitably isotopically labelled precursors can be used to generate isotopomeric mixtures to allow flux studies when used in combination with isotopomers of nicotinamide. While D₄-labelled Nam is commercially available, labelling of Nam with ¹⁸O proved to provide good traceability characteristics as it offers a stable MW+2 profile in the NAD⁺ metabolites which incorporate it²¹. In order to enable the scalable syntheses of such isotope containing species, improved methods to access ¹⁸O-Nam (7) were required (Scheme 3).

To generate large quantities of ¹⁸O-Nam (7) with minimum cost, minimizing the amount of ¹⁸OH₂ used in this reaction was paramount, and thus both mechanochemistry and sealed reaction conditions were explored. Best conversions were obtained when sealed conditions and 3.3-fold molar excess of ¹⁸OH₂ (Cambridge Isotope Laboratory; 97% ¹⁸O incorporation) were used in the hydrolysis of cyanopyridine in presence of manganese oxide. The use of U-¹³C-labelled tetraacetate riboside (9) combined to the ¹⁸O-labelled Nam, allowed for the generation of MW+7 NR (11)⁴² and NRH (13) isotopologues (overall 63% and 60% yields from [¹³O₅]ribose for (MW+7) NR-OTf and (MW+7) NRH, respectively). Isotopic incorporation in 11 and 13 was determined by high resolution MS with the MW+7 species accounting for more than 94%, in both cases.

The ¹⁸O labelling of NAR and NARH is unsuitable for metabolomic measurements as 50% of the label is lost upon metabolic conversion to NAD⁺. However, the ¹³C-nitrile (14) generated from bromo-pyridine and K¹³CN offers an alternative labelling pattern. With this precursor, isotopologues of Nam (MW+3) (15) with 96% isotopic incorporation and NA (MW+1) (16) with isotopic incorporation of ca. 100% (Scheme 4) were generated at scale suitable for animal studies and at relatively low cost. Overall, we have developed easily implementable and relatively low-cost methods to access isotopically labelled derivatives of vitamin B3.

Conclusions

We have combined mechanochemistry and standard solution phase synthesis to optimize the synthesis of NR, NRH, NAR and NARH with the view of improving synthetic efficiencies and enabling scale-ups, as well as facilitating the incorporation of isotopic labels in these scaffolds. Chemical stability differences between the acid and amide versions of these ribosylated species have also been highlighted as they impact synthetic efficacy. These NAD⁺ precursors and their isotopologues give insights on yet unexplored modes of action of NAD⁺ and how it can be modulated.

Supplementary Material