Bioavailability studies indicated that ingested NAD+ was primarily hydrolyzed in the small intestine by brush border cells (Baum et al., 1982, Gross and Henderson, 1983). As a first step, NAD+ is cleaved to NMN and 5′-AMP by a pyrophosphatase found either in intestinal secretions (Gross and Henderson, 1983) or in the brush border (Baum et al., 1982). Next, NMN is rapidly hydrolyzed to NR, which in turn is more slowly converted into NAM (Gross and Henderson, 1983). NAM can also be formed directly by the cleavage of NAD+, obtaining ADP-ribose derivates as a side product (Gross and Henderson, 1983). The intestinal production of NAM from NAD+ or NR required the presence of intestinal cells, indicating that the enzymes for this process are membrane-bound or intracellular (Baum et al., 1982, Gross and Henderson, 1983). The direct perfusion with NAM, however, did not give rise to any of these species, indicating that NAM is the final degradation product and directly absorbed (Collins and Chaykin, 1972, Gross and Henderson, 1983, Henderson and Gross, 1979). In contrast, perfusion of the intestine with NA revealed a substantial cellular accumulation of labeled intermediates of the NAD+ biosynthetic pathway, including NAM, which suggest the presence of active NA metabolism in intestinal cells (Collins and Chaykin, 1972, Henderson and Gross, 1979). In line with this, blood concentrations of NA are relatively low (∼100 nM) yet, when pharmacologically primed (Jacobson et al., 1995, Tunaru et al., 2003), can increase and be rapidly converted to NAM by the liver (Collins and Chaykin, 1972). Strikingly, NAM levels in fasted human plasma are also too low to support NAD+ biosynthesis in cells (between 0.3 and 4 μM) (Hara et al., 2011, Jacobson et al., 1995). All of these results suggest that these NAD+ precursors are metabolized very quickly in mammalian blood and tissues.
Lipid-Lowering Effect of Niacin
NA attracted clinical attention for its cholesterol lowering actions (Altschul et al., 1955), and became the first drug used to treat dyslipidemia. Gram dosages of NA reduce plasma triglyceride and low-density lipoprotein (LDL) levels, while concomitantly increasing high-densitiy lipoproteins (HDL). However, the clinical use of NA has been limited by the fact that it induces cutaneous flushing, which compromises compliance (Birjmohun et al., 2005). This flushing does not derive from the ability of NA to drive NAD+ synthesis, but rather from the activation of a G protein-coupled receptor, GPR109A (Benyó et al., 2005). Given the low presence of NA in blood, the activation of this receptor is unlikely to be a native function of NA, but rather an effect from pharmacological dosing. It was also assumed that the beneficial effects of NA on plasma lipids are mediated via a receptor rather than a vitamin mechanism because of the high dose required (100-fold higher than that required to prevent pellagra) and the failure of NAM to provide similar benefits (Tunaru et al., 2003). Indeed, some evidence supports that GPR109A is necessary for NA to raise HDL cholesterol (Li et al., 2010, Tunaru et al., 2003). However, the absence of GRP109A expression in the liver (Soga et al., 2003, Tunaru et al., 2003, Wise et al., 2003), a central hub for HDL and LDL metabolism, also questions whether the effects of NA on blood lipids derive from GPR109A activation. Alternatively, strong evidence for the ability of NAD+ to enhance the activity of sirtuins provides a mechanism of action that also drives benefits on lipid homeostasis (Cantó and Auwerx, 2012). In addition, sirtuin activity is inhibited by NAM (Anderson et al., 2003), which could explain why NAM failed to provide the benefits of NA; however, in some situations, NAM treatments can have beneficial effects as discussed in New Perspectives in NAD+ Therapeutics (I): Metabolic Disease. The intricate relationship between NAD+ and sirtuins will be discussed further in NAD+-Consuming Enzymes (I): Sirtuins.
Introducing NAD+ as a Metabolic Regulator
The role of NAD+ as a coenzyme in most metabolic pathways suggests that NAD+ limitations could affect metabolic efficiency. Decreasing NAD+ levels could therefore prompt the development of many of the ailments associated with aging. Indeed, NAD+ levels can change during a number of physiological processes. Diverse lines of research on worms, rodents, and human cellular models indicate that declining NAD+ levels are a hallmark for senescence (Braidy et al., 2011, Gomes et al., 2013, Khan et al., 2014, Massudi et al., 2012, Mouchiroud et al., 2013, Ramsey et al., 2008, Yoshino et al., 2011). Along a similar line, a reduction in muscle progenitor cell NAD+ content leads to a SIRT1-mediated metabolic switch that induces premature differentiation and a loss of regenerative capacity, reflecting a phenotype typical of aging muscle (Ryall et al., 2015). The link between metabolism and NAD+ is further solidified by observations that tissue NAD+ levels decrease with high-fat diets (HFDs) (Bai et al., 2011b, Cantó et al., 2012, Kraus et al., 2014, Pirinen et al., 2014, Yang et al., 2014, Yoshino et al., 2011). In contrast, NAD+ increases in mammalian cells and tissues in response to exercise (Cantó et al., 2009, Cantó et al., 2010, Costford et al., 2010) or calorie restriction (CR) (Chen et al., 2008), both of which are interventions associated with metabolic and age-related health benefits. In line with this, supplementation with NAD+ precursors has proven to enhance lifespan in budding yeast (Belenky et al., 2007) and worms (Mouchiroud et al., 2013). Also, in mammals, the enhancement of NAD+ levels has been linked with improved mitochondrial function under stress (Cerutti et al., 2014, Khan et al., 2014, Mouchiroud et al., 2013, Pirinen et al., 2014), leading to protection against dietary (Bai et al., 2011b, Cantó et al., 2012) and age-related (Gomes et al., 2013, Yoshino et al., 2011) metabolic complications. Finally, hepatic NAD+ levels dynamically change in a circadian fashion (Asher et al., 2010, Nakahata et al., 2009, Ramsey et al., 2009), weaving an intricate relationship with nutritional states. Therefore, despite the classical misconception that intracellular NAD+ levels rarely change (Kaelin and McKnight, 2013), the evidence above unequivocally demonstrates the ability of NAD+ to respond dynamically to physiological stimuli. So, how do changes in NAD+ levels take place innately?
NAD+ Synthesis and Salvage: New Ways to Boost NAD+
NAD+ Biosynthesis and the Discovery of New NAD+ Precursors
NAD+ availability is determined by the relative rates of NAD+ biosynthesis and degradation. Ergo, the enhancement of NAD+ biosynthesis could provide a way to elevate NAD+ content. There are several known NAD+ precursors. First, dietary Trp can serve as an NAD+ precursor through an eight-step de novo pathway (Bender, 1983), which has been described in detail elsewhere (Houtkooper et al., 2010a); so, we will only focus on some of its most interesting features (Figures 1A–1D). The first and rate-limiting step in this path includes the conversion of Trp to N-formylkynurenine by either indoleamine 2,3-dioxygenase (IDO) or tryptophan 2,3-dioxygenase (TDO) (Figure 1B). These enzymes are strongly overexpressed in diverse cancers, and the subsequent synthesis of kynurenines may act as potential second messengers in cancer immune tolerance (Stone and Darlington, 2002), possibly through binding to the aryl hydrocarbon receptor (AHR) (Bessede et al., 2014). An interesting branch point in the Trp catabolic pathway is the formation of the unstable α-amino-β-carboxymuconate-ε-semialdehyde (ACMS) (Bender, 1983). ACMS can be enzymatically converted to α-amino-β-muconate-ε-semialdehyde (AMS) by ACMS decarboxylase (ACMSD), leading to complete oxidation via the glutarate pathway and the tricarboxylic acid (TCA) cycle or to the production of picolinic acid via a spontaneous reaction (Figures 1B and 1C) (Houtkooper et al., 2010a). Alternatively, ACMS can undergo spontaneous cyclization forming quinolinic acid, which subsequently serves as an NAD+ precursor (Bender, 1983). This latter nonenzymatic possibility seems to be only relevant when the metabolism of ACMS is limited in the cell. This might explain why, in general, Trp is considered a rather poor NAD+ precursor in vivo, as it will only be diverted to NAD+ synthesis when its supply exceeds the enzymatic capacity of ACMSD (Ikeda et al., 1965). In humans, diets ranging from 34 mg to 86 mg of Trp provide the equivalent of 1 mg of niacin (reviewed in Horwitt et al., 1981). Interestingly, the formation of NAD+ following Trp injections is further reduced in diabetic rats (Ikeda et al., 1965). When ACMSD capacity is surpassed, Trp-derived quinolinic acid is produced and used by quinolinate phosphoribosyltransferase (QPRT) to form NA mononucleotide (NAMN). NAMN is then converted to NA adenine dinucleotide (NAAD), using ATP, by the enzyme NMN adenylyltransferase (NMNAT) (Figure 1A) (Houtkooper et al., 2010a). This is a key enzyme for NAD+ synthesis in mammals, irrespective of the precursor used, since it is also needed for NAD+ salvage. Three NMNAT isoforms (NMNAT1–3) with different tissue and subcellular distributions have been described in mammals (Lau et al., 2009). NMNAT1 is a nuclear enzyme that is ubiquitously expressed, with its highest levels in skeletal muscle, heart, kidney, liver, and pancreas, yet it is almost undetectable in the brain (Emanuelli et al., 2001). In contrast, NMNAT2 is mostly located in the cytosol and Golgi apparatus (Berger et al., 2005, Yalowitz et al., 2004). Finally, NMNAT3 is highly expressed in erythrocytes with a moderate expression in skeletal muscle and heart and has been identified in both cytosolic and mitochondrial compartments, with cell-/tissue-specific subcellular localization patterns (Berger et al., 2005, Felici et al., 2013, Hikosaka et al., 2014, Zhang et al., 2003). The possible implications of the subcellular localization of NMNAT enzymes will be discussed in Cell Compartmentalization of NAD+. The last step in the primary biosynthesis of NAD+ includes the ATP-dependent amidation of NAAD by NAD+ synthase (NADSYN) using glutamine as a donor. NADSYN is mainly expressed in the small intestine, liver, kidney, and testis, where this pathway may be more relevant to NAD+ synthesis (Hara et al., 2003, Houtkooper et al., 2010a).
NAD+ can also be synthesized from metabolite recycling or the dietary uptake of other NAD+ precursors (Houtkooper et al., 2010a). NA can lead to NAD+ through the shorter, three-step Preiss-Handler pathway (Figure 1A). Here, NA is initially metabolized by the NA phosphoribosyltransferase (NAPRT) into NAMN, converging with the de novo pathway.
In mammals, NAM can also be an NAD+ precursor through its metabolism into NAM mononucleotide (NMN) by the rate-limiting enzyme nicotinamide phosphoribosyltransferase (NAMPT) (Figure 1D) (Revollo et al., 2004, Rongvaux et al., 2002). NMN can be then converted into NAD+ through a single additional reaction catalyzed by the NMNAT enzymes. NAM is also the product of NAD+ degradation by several enzyme families (see The Enzymatic Use of NAD+). Consequently, NAMPT is key to not only metabolizing circulating NAM, but also to recycling intracellularly produced NAM via the NAD+ salvage pathway. As a key enzyme, SNPs found in non-coding regions of human NAMPT are correlated with glucose and lipid metabolism alterations and type 2 diabetes, among other disease associations (Zhang et al., 2011).
Lastly, NR metabolism constitutes an additional path for NAD+ biosynthesis (Bieganowski and Brenner, 2004) (Figure 1D). NR is transported into cells by nucleoside transporters (Nikiforov et al., 2011) and is then phosphorylated by the NR kinases 1 and 2 (NRKs) (Bieganowski and Brenner, 2004), generating NMN. This phosphorylation step is a conserved feature in all eukaryotes (Bieganowski and Brenner, 2004), underscoring its evolutionary relevance. After the generation of NMN, NMNAT enzymes can then catalyze the formation of NAD+. While additional ways for NR metabolism have been described in yeast (Belenky et al., 2007), the phosphorylation by NRKs is still the only pathway described in mammalian cells for the transformation of NR into NAD+.
Whole-Body NAD+ Transport
Despite Trp being the canonical NAD+ precursor, its action may be up to 60 times less efficient than NA (Institute of Medicine (US) Standing Committee on the Scientific Evaluation of Dietary Reference Intakes and its Panel on Folate, Other B Vitamins, and Choline, 1998), as Trp is also used for protein translation and other biosynthetic purposes. Indeed, the use of Trp as an NAD+ precursor would not be solely sufficient to support the physiological NAD+ requirements in mammals (Henderson, 1997). NA, in contrast, can act as a potent NAD+ precursor, primarily in liver and kidney where NAPRT demonstrates the highest activity levels (Hara et al., 2007). However, mammalian tissues are rarely exposed to NA, as its levels in blood are generally very low (Jacobson et al., 1995, Tunaru et al., 2003). Also, as discussed in Food Sources and Bioavailability of NAD+, most evidence suggests that NA might be quickly metabolized to NAM in the gut and the liver (Collins and Chaykin, 1972). However, the low plasma concentration of NAM (Hara et al., 2011) is ∼1,000-fold less than that required to increase NAD+ levels in cultured cells (Hara et al., 2007, Revollo et al., 2004).