Journal of Experimental Botany, Vol. 57, No. 8, pp. 1603–1620, 2006
Oxygen Metabolism, ROS and Redox Signalling in Plants Special Issue
doi:10.1093/jxb/erj202 Advance Access publication 12 May, 2006
NAD(P) synthesis and pyridine nucleotide cycling in plants
and their potential importance in stress conditions*
Graham Noctor†, Guillaume Queval and Bertrand Gakière
Institut de Biotechnologie des Plantes, Université de Paris XI, F-91405 Orsay, France
Received 1 March 2006; Accepted 17 March 2006
Abstract
Introduction
Pyridine nucleotides are key redox carriers in the
soluble phase of all living cells, and both NAD and
NADP play crucial roles in pro-oxidant and antioxidant
metabolism. Recent data also suggest a number of
non-redox mechanisms by which these nucleotides
could influence cell function. In cases where these
mechanisms involve NAD(P) consumption, resynthesis
must occur to maintain nucleotide pools. Important information on the pathways involved in NAD synthesis
in plants is beginning to appear, but many outstanding
questions remain. This work provides an overview of
the current state of knowledge on NAD synthesis pathways in plants and other organisms, analyses plant
sequences for the first two enzymes of the de novo
synthesis of NAD, proposes a preliminary model for
the intracellular distribution of NAD synthesis, presents plant homologues of recently identified yeast
mitochondrial NAD transporters, and discusses factors
likely to be important in the regulation of NAD synthesis and contents in plants, with particular reference to
stress conditions.
NAD and its derivative NADP are ubiquitous in living cells
(Penfound and Foster, 1996; Kurnasov et al., 2003). They
mediate hundreds of redox reactions, and thus impact on
virtually every metabolic pathway in the cell. Pyridine
nucleotides are also key players in signalling through
reactive oxygen species (ROS) since they are crucial in
the regulation of both ROS-producing and ROS-consuming
systems in plants (Møller, 2001; Apel and Hirt, 2004;
Mittler et al., 2004; Foyer and Noctor, 2005). Chloroplast
ROS production is influenced by NADP+:NADPH ratios,
mitochondrial NAD(P) status is important in determining
ROS formation by the respiratory electron transport chain,
and NAD(P)H oxidases are key players in the generation
of ROS at the plasmalemma. Similarly, ROS processing
partly depends on ascorbate and glutathione, and redox
flux through these pools is maintained by NAD(P)H.
Pyridine nucleotides are involved in many other defence
and signalling reactions, for example, production of nitric
oxide and metabolism of reactive lipid derivatives (Crawford
and Guo, 2005; Mano et al., 2005). Thus, as well as their
roles as cofactors in numerous energy-producing and biosynthetic reactions, NAD and NADP are crucial in redox
signalling linked to stress and development.
Recent studies show that NAD and NADP play important roles in cell signalling in animals, plants, and fungi
(Hunt et al., 2004; Ziegler, 2005). NAD and NADP are the
respective substrates for the production of the calcium
agonists cyclic ADP-ribose (cADP-R; Sánchez et al., 2004)
and nicotinate adenine dinucleotide phosphate (NaADP;
Navazio et al., 2000; Lee, 2005). Transfer of ADP-ribose
Key words: L-Aspartate oxidase, compartmentation, NAD transporter, NAD signalling, nicotinamide, nicotinate, quinolinate
synthase.
* In this article, NAD(P) is used to denote total pools of reduced and oxidized forms or in cases where the distinction between the two forms is unnecessary
(e.g. NAD synthesis). NAD(P)+ is used to make specific reference to the oxidized forms.
y
To whom correspondence should be addressed. E-mail: [email protected]
Abbreviations: (c)ADP-R, (cyclic)ADP-ribose; CMS, cytoplasmic male sterile; DHAP, dihydroxyacetone phosphate; MNam, N-methyl nicotinamide; NaAD,
nicotinate adenine dinucleotide; NaADP, nicotinate adenine dinucleotide phosphate; NADase, NAD glycohydrolase; NaMN(AT), nicotinate mononucleotide
(adenylyl transferase); NaPRT, nicotinate phosphoribosyltransferase; NMDA, N-methyl-D-aspartate; NMN(AT), nicotinamide mononucleotide (adenylyl
transferase); NRK, nicotinamide riboside kinase; PARP, poly(ADP-ribose) polymerase; QPRT, quinolinate phosphoribosyltransferase; ROS, reactive oxygen
species; SIRTUIN, homologue of the yeast Silent Information Regulator 2 protein.
ª The Author [2006]. Published by Oxford University Press [on behalf of the Society for Experimental Biology]. All rights reserved.
For Permissions, please e-mail: [email protected]
1604 Noctor et al.
units from NAD to target proteins is an important posttranslational modification, catalysed by poly(ADP-ribose)
polymerases (PARPs) and mono(ADP-ribosyl)transferases
(Ying et al., 2003; Graziani et al., 2005). The yeast Silent
Information Regulator 2 (Sir2) protein has been shown to
be an NAD-dependent deacetylase (Imai et al., 2000; Lin
et al., 2000; Berger et al., 2004; Ziegler, 2005). Although
these components and phenomena are better characterized
in other systems than in plants, many of the products and
the enzymes that produce them are also found in plants
(Hunt et al., 2004). Since some of these NAD(P)-consuming
reactions are important in stress conditions and/or are stress
inducible, NAD concentration could be influential in plant
responses to environmental challenge. Exposure of plant
cells to oxidative stress produces decreases in overall
NAD contents, linked to PARP activity (Amor et al.,
1998; De Block et al., 2005). Recent studies of mutants
provide evidence that changes in NAD status can alter
photosynthesis and plant stress responses (Dutilleul et al.,
2003a, b; Hayashi et al., 2005), and suggest that NAD
content could be a powerful modulator of metabolic
integration (Dutilleul et al., 2005). Much remains to be
discovered about the factors controlling pyridine nucleotide synthesis and recycling in plants.
NAD recycling and associated reactions
The most studied route to NAD in mammals is via
nicotinamide and nicotinate. Together, these compounds
are called ‘niacins’ and were identified by their ability to
prevent pellagra in dogs fed a diet poor in tryptophan, which
is a precursor in the de novo synthesis of the pyridine ring in
animals (see below). It was subsequently shown that niacins
can be converted to NAD via nicotinate phosphoribosyltransferase (NaPRT), nicotinate mononucleotide adenylyltransferase (NaMNAT), and NAD synthetase (Preiss and
Handler, 1958). Recently, the structure of a yeast NaPRT
has been determined at a resolution of 1.75 Å (Chappie
et al., 2005). In animals, which lack nicotinamide deamidase activity (Chappie et al., 2005), NAD can also be
produced via nicotinamide phosphoribosyltransferase
(NPRT) activity, followed by adenylylation of NMN.
This allows incorporation of nicotinamide into NAD in
two steps rather than the four-step pathway via nicotinate
(Fig. 1, orange background).
Labelling studies in plants suggest that nicotinamide
released from NAD is first deamidated by nicotinamidases
to nicotinate (Ryrie and Scott, 1969; Ashihara et al., 2005;
Zheng et al., 2005) which is then converted to NaMN by
NaPRT (Fig. 1). This view is supported by the presence in
Arabidopsis and other plants of genes predicted to encode
NaPRT, NaMNAT, and NAD synthetase (Hunt et al.,
2004; Katoh and Hashimoto, 2004). Consistent with this,
homologues of mammalian NPRT gene sequences appear
to be absent from plants (Katoh and Hashimoto, 2004) and
NPRT activity has not been detected in extracts of tobacco,
mungbean, or white spruce cells (Wagner et al., 1986;
Ashihara et al., 2005; Zheng et al., 2005).
Results of a detailed study of the fate of 14C-labelled
nicotinamide in coffee and cocao (Zheng et al., 2004) also
suggest that nicotinate is the main precursor of pyridine
nucleotides, although NMN was also labelled in significant amounts, probably as a breakdown product of NAD
through NAD pyrophosphatase activity (Fig. 1). In view of
these results, together with data gathered from labelling
studies in white spruce and mungbean (Ashihara et al.,
2005; Zheng et al., 2005), three possible routes were
suggested from NAD to nicotinamide. The first involves
production of nicotinamide in a single step, with ADP-R
(or derivatives) as the other product. This route would
result from the activities of enzymes such as NAD glycohydrolase, PARP, SIRTUINS, and ADP-ribosyl cyclase
(Fig. 1). A second pathway would proceed via NMN
and involve two steps, catalysed by NAD pyrophosphatase
and NMN nucleosidase (Zheng et al., 2004). In the third
pathway, NMN is indirectly converted to nicotinamide via
nicotinamide riboside, and overall the three steps involve
NAD pyrophosphatase, 5’ nucleotidase, and nicotinamide
nucleoside phosphorylase or nucleosidase (Ashihara et al.,
2005; Zheng et al., 2005). According to this scenario,
full turnover of the pyridine cycling pathway (from NAD
to NAD) could involve five, six, or seven steps. The final
four steps would be common to all these recycling
pathways and involve formation of nicotinate from nicotinamide, conversion of nicotinate to NaMN, adenylylation
of NaMN to NaAD (desamidoNAD), and amidation of
NaAD to produce NAD (Fig. 1).
Given the presence of NMN in plant tissues, further work
is required to confirm the substrate specificities of plant
N(a)MNAT activities, and to establish whether these enzymes are specific to NaMN. In mammals, three adenylyltransferases have been characterized and all are able to use
NaMN and NMN (Magni et al., 2004). This probably
reflects the dual role of mammalian N(a)MNAT in both
the de novo pathway (production of NaAD from NaMN)
and the recycling pathway (production of NAD from
NMN; Fig. 1, orange background, top). In bacteria, the
adenylyltransferase shows a clear preference for NaMN, a
compound which in these organisms is common to both
de novo synthesis and the recycling pathway from nicotinamide by the successive action of the pncA and pncB gene
products (Fig. 1). The final step of NAD synthesis is catalysed by NAD synthetases, probably using glutamine,
although in some organisms isoforms are able to use ammonia under physiological conditions (Magni et al., 2004).
In addition to synthesis of pyridine mononucleotides
(NMN and NaMN) by phosphoribosyltransferase activities,
a third path to NAD+ has been described in species of
bacteria, yeast, and mammals (Bieganowski and Brenner,
2004). This route involves synthesis of NMN by
NAD(P) synthesis and pyridine nucleotide cycling
phosphorylation of nicotinamide riboside (Fig. 1). An
elegant vitamin screen using yeast mutants indicated the
presence of nicotinamide riboside in milk (Bieganowski
and Brenner, 2004). As discussed above, nicotinamide
riboside might be formed in mungbean from NMN (Zheng
et al., 2005). A BLAST search of several plant genome
databases using bacterial, yeast, and mammalian sequences
for nicotinamide riboside kinase revealed no fully homologous sequences, although homologous domains can be
identified among the uridine kinase family.
In addition to nicotinamide riboside, plants contain
nicotinate riboside. Indeed, labelling studies in coffee
have provided some evidence that this compound is an
intermediate in pyridine cycling and that NaMN can be
formed from nicotinate through two paths (Zheng et al.,
2004). As well as direct phosphoribose transfer catalysed
by NaPRT, a second path is possible through successive
ribosylation and phosphorylation (Fig. 1, orange background, bottom). These reactions are reversible and could
also represent a route from NaMN to nicotinate (Ashihara
et al., 2005). If they operate in the direction of NaMN
synthesis, it would add yet another step to the recycling
pathway in plants. In this case, recycling of the pyridine
ring from NAD to NAD could potentially consist of eight
steps rather than the five-step pathway described in bacteria
(Fig. 1, orange background, cf. shorter route traced by the
solid arrows with the longer path including dotted arrows
via the two ribosides).
Methylation of nicotinamide and nicotinate produces Nmethyl nicotinamide (MNam) and N-methyl nicotinate
(trigonelline). In mammals, MNam is the initial product
of a degradation pathway in which pyridone products are
subsequently formed by MNam oxidation (Fig. 1, grey
background). Urinary excretion of both MNam and pyridone products is stimulated when the diet is rich in
nicotinamide or nicotinate (Creeke and Seal, 2005, and
references therein). MNam has been detected in plants
(Waller et al., 1966) while trigonelline is known to be
formed in many types of plants, notably legumes, and
may have hormonal-type functions and be important as a
store of nicotinate (Minorsky, 2002; Zheng et al., 2004,
2005). In addition to MNam and trigonelline, other plant
compounds could act as conjugated stores of nicotinamide and nicotinate, including N-glucosylated forms
and nicotinate riboside (Berglund et al., 1996; Zheng
et al., 2004). In coffee and cocoa, most of the label
from [14C]nicotinamide was incorporated into trigonelline,
with only minor labelling of nicotinate riboside and
nicotinate glucoside (Zheng et al., 2004). In contrast, in
tea, the major labelled compound was nicotinate glucoside,
with no detectable label found in trigonelline. These
observations show that the type of nicotinate conjugate
formed is species dependent (Zheng et al., 2004).
As well as trigonelline, other pyridine-based alkaloids
include ricinine, nicotine, and anabasine (Fig. 1, blue
1605
background). Production of these compounds is enhanced
by stress, particularly wounding, and they are effective
feeding deterrents. Much of the information that has been
gathered on the regulation of NAD-linked metabolism in
plants has come from investigations of pyridine alkaloid
synthesis rather than NAD(P) synthesis per se (Waller
et al., 1966; Frost et al., 1967; Mann and Byerrum, 1974a,
b; Wagner et al., 1986; Sinclair et al., 2000; Cane et al.,
2004; Zheng et al., 2004, 2005). Labelling studies have
shown that nicotinate, nicotinamide, quinolinate, NaAD,
and NAD are all effective sources of the pyridine ring for
nicotine and ricinine synthesis (Waller et al., 1966; Frost
et al., 1967). These data emphasize the close relationship
between NAD metabolism and alkaloid production in these
species. The observation that quinolinate and NAD are as
effective as nicotinate in supporting production of pyridine
alkaloids (Waller et al., 1966; Frost et al., 1967; Zheng
et al., 2005) suggests that, at least in those species
examined, the pyridine recycling pathway is capable of
relatively rapid turnover under many conditions.
NADP(H) synthesis and utilization
NADP is produced from NAD by NAD(H) kinase. At least
one NAD kinase in tobacco is a calmodulin-binding
protein (Harding et al., 1997) and activity in wild tomato
may be subject to regulation by thiol/disulphide exchange
(Delumeau et al., 2000). NAD kinase activity could be
located in the chloroplast, cytosol, and mitochondria
(Gallais et al., 2000a; Turner et al., 2004; Chai et al.,
2005). Two NAD+-specific Arabidopsis NAD kinases,
named NADK1 and NADK2, have been cloned and
characterized biochemically (Turner et al., 2004, 2005).
Compared with NADK1, NADK2 has a large N-terminal
extension (Turner et al., 2004), probably involved in regulation by calmodulin (Harding et al., 1997). Total extractable NAD kinase was shown to be highly dependent
on calmodulin in Arabidopsis roots and shoots throughout
development (Turner et al., 2004). In contrast to NADK1
and NADK2, a third characterized Arabidopsis isoform,
NADK3, shows a strong preference for NADH over NAD
(60-fold lower Km for NADH; Turner et al., 2005). The
yeast homologue of Arabidopsis NADK3 is a mitochondrial
enzyme, whereas the plant enzyme appears to be located
in the cytosol (Turner et al., 2005). In plants, mitochondrial NADP could be at least partly imported from the
cytosol (Bykova and Møller, 2001). If NADK3 is a cytosolic enzyme, some in vivo activity with NAD+ cannot be
completely discounted in view of the high total NAD+/
NADH ratio in this compartment (10–30: Igamberdiev and
Gardeström, 2003). Indeed, the physiologically relevant
‘free’ NAD+/NADH ratio (nucleotides not bound to protein
at any given moment) may be as high as 600–1000 in the
cytosol of both plant and mammalian cells (Heineke et al.,
1991; Zhang et al., 2002). Other authors have reported that
1606 Noctor et al.
Fig. 1. Principal reactions in NAD synthesis and metabolism. The scheme incorporates elements from current information on both plants and other
organisms. The solid arrows show the de novo synthesis pathway and the five-step NAD cycling pathway established in bacteria. In addition, a three-step
recycling pathway through NMN operates in animals, NAD can be synthesized from nicotinamide riboside in several organisms (Bieganowski and
Brenner, 2004), and it has been proposed that the cycling route in some plants could involve six, seven, or even eight steps (Zheng et al., 2004, 2005).
Reactions necessary for these pathways are shown as dotted arrows on the orange background. Some important pyridine alkaloids are shown against the
blue background. Dashed arrows show other routes to quinolinate apparently not present in plants. Bacterial nad [NAD(P) synthesis] and pnc (pyridine
nucleotide cycle) homologue names are indicated in parentheses for some of the enzymes. Abbreviations: AdoHcys, S-adenosylhomocysteine; AdoMet,
NAD(P) synthesis and pyridine nucleotide cycling
Arabidopsis NADK2 appears to localize to the chloroplast and that an insertional mutant for the enzyme shows
stunted growth and stress hypersensitivity, associated with
a minor decrease in total tissue NADPH (Chai et al., 2005).
NADP+ phosphatase activity, which converts NADP+ to
NAD+, has also been reported in plants (Gallais et al., 2000b),
and the mitochondrial NAD(H) and NADP(H) pools are
redox-linked via transhydrogenases (Bykova et al., 1999).
As well as its redox cofactor function, NADP is the
precursor for the calcium channel activator NaADP (Lee,
2005). NaADP has been shown to be active in plants
(Navazio et al., 2000) and can be synthesized through base
exchange of the nicotinamide moiety of NADP with free
nicotinate, a reaction catalysed by ADP-ribosyl cyclases
under certain conditions (Fig. 1, green background, left).
NaADP concentrations could also be regulated by specific
calcium-dependent phosphatases, which convert it to
NaAD (Berridge et al., 2002), or nucleotide pyrophosphatases, which produce NaMN and AMP-29-phosphate
(Evans et al., 2005; Fig. 1). Glycohydrolases can function
to degrade NADP to nicotinamide and ADP-R 29phosphate or cADP-R 29-phosphate. The best-characterized
ADP-ribosyl cyclase in mammals is the membrane-bound
glycoprotein CD38 (Evans et al., 2005). This protein can
convert extracellular or intracellular NAD to cADP-R,
hydrolyse cADP-R to ADP-R, and catalyse synthesis of
NaADP via the base exchange reaction with nicotinate
(Fig. 1, green and grey backgrounds). Although no genes
have been identified on the basis of sequence homology,
low activities of both ADP-ribosyl cyclase and cADP-R
hydrolase have been detected in Arabidopsis, and cADP-R
has been shown to accumulate transiently in response to
abscisic acid (Sánchez et al., 2004). In human platelets, it
has also been reported that CD38 can cleave NADP to
ADP-R 29-phosphate (Torti et al., 1999). Like the reactions
catalysed by PARP and SIRTUINS, many of these NAD
and NADP cleavage reactions produce nicotinamide
(Fig. 1, green background).
De novo synthesis of nicotinate mononucleotide
(NaMN)
As well as producing NaMN from nicotinate obtained from
the environment or formed during the recycling pathway,
almost all organisms appear to be able to produce this NAD
1607
precursor via synthesis of the pyridine ring in the form of
quinolinate (pyridine 2,3-dicarboxylate; Fig. 1, purple
background). NaMN is then produced by quinolinate
phosphoribosyltransferase (QPRT; EC 2.4.2.19), encoded
by nadC in Escherichia coli (Fig. 1). Characterization of
the enzyme purified from castor bean endosperm showed
that Km values for quinolinate and 5-phosphoribosyl-1pyrophosphate were 12 and 45 lM, respectively (Mann and
Byerrum, 1974a). Tobacco QPRT cDNA sequences
suggested that the protein is probably located in the
mitochondria, although a shorter transcript might produce
a cytosol-directed protein (Sinclair et al., 2000). Computer
analysis (http://urgi.infobiogen.fr/predotar) of sequences
encoded by the single QPRT gene in Arabidopsis (Fig. 1)
predicts two transcripts, the longer of which probably
encodes a mitochondrial import sequence. Quinolinatedependent synthesis of NaMN therefore probably occurs in
both cytosolic and mitochondrial compartments.
In animals and fungi, quinolinate is formed through
a multistep pathway from tryptophan. Recently, gene
sequences for several enzymes of this pathway have also
been found in certain pathogenic bacteria (Kurnasov et al.,
2003), and some anaerobic species have been reported to
make quinolinate by acetylation of N-formyl-L-aspartate
(Scott et al., 1969; Griffith et al., 1975). However, by far
the best characterized prokaryotic pathway for quinolinate
synthesis is from aspartate and triose phosphate (Griffith
et al., 1975; Flachmann et al., 1988; Kurnasov et al., 2003).
This pathway involves two enzymatic steps, catalysed by
quinolinate synthase, a term which has been used to refer
to the two enzymes functioning together or specifically to
designate the second enzyme. The first enzyme, L-aspartate
oxidase (or quinolinate synthase ‘B’), is encoded by nadB,
while the second, quinolinate synthase (or quinolinate
synthase ‘A’), is encoded by nadA (Fig. 1). Close association of the two enzymes might enable the intermediate,
iminoaspartate, an unstable compound that spontaneously
hydrolyses to oxaloacetate, to be channelled from the first
enzyme to the second. In E. coli, the nadB and nadA genes
are arranged in a single operon together with the nadC
gene encoding QPRT (Griffith et al., 1975).
Quinolinate synthesis in plants
In mammals, in addition to the well-known neuroactivity
of nicotine, the endogenously produced NAD precursor
S-adenosylmethionine; ADP-R cyc, ADP-ribosyl cyclase; (c)ADP-R, (cyclic)ADP-ribose; 29-P-(c)ADP-R, (cyclic)ADP-ribose-29-phosphate; 29-PAMP, AMP-29-phosphate; DHAP, dihydroxyacetone phosphate; MNam, N1-methylnicotinamide; MRT, mono(ADP-ribose)transferase; NaAD,
nicotinate adenine dinucleotide; NaADP, nicotinate adenine dinucleotide phosphate; NADase, NAD glycohydrolase (NAD nucleosidase); NAD ppase,
NAD pyrophosphatase; NADP+ pase, NADP+ phosphatase; NAD-S, NAD synthetase; NaMN(AT), nicotinate mononucleotide (adenylyltransferase);
NaPRT, nicotinate phosphoribosyltransferase; NaRK, nicotinate riboside kinase; 59Nase, 59-nucleotidase; NMN(AT), nicotinamide mononucleotide
(adenylyltransferase); NMN nase, NMN nucleosidase; (N)NP, (nicotinamide) nucleoside phosphorylase; N ppase, nucleotide pyrophosphatase; NRK,
nicotinamide riboside kinase; PARP, poly(ADP-ribose)polymerase; 29 Pase, 29 phosphatase; Pi, phosphate; Ppi, pyrophosphate; Prot, protein; PRPP, 59phosphoribosyl-19-pyrophosphate; 2-PYR, N-methyl-2-pyridone-5-carboxamide; 4-PYR, N-methyl-4-pyridone-3-carboxamide; QPRT, quinolinate
phosphoribosyltransferase; R1P, ribose 1-phosphate; SIRTUINS, homologues of the yeast Silent Information Regulator 2 (Sir2) protein. Not all possible
interconversions are shown. Double-headed arrows indicate some of the reactions that are theoretically reversible.
1608 Noctor et al.
quinolinate acts as an agonist at N-methyl-D-aspartate
(NMDA) receptors, which are classed among the glutamate
receptor family (Stone and Addae, 2002). Quinolinate
accumulation in the brain has been implicated in several
neurodegenerative disorders (Fukuoka et al., 2002). The
Arabidopsis genome contains numerous loci that encode
subunits of glutamate receptors. Although much remains to
be discovered concerning the functions of these proteins,
plant NMDA receptor-like proteins share certain characteristics of their animal counterparts (Dubos et al., 2003).
It is an intriguing possibility that, in addition to NAD(P)linked signalling functions associated with redox control,
production of NaADP and cADP-ribose, and PARP and
SIRTUIN activities, other NAD(P) precursors or metabolites such as quinolinate might act as information transmitters in plants.
To date, there have been very few studies of the enzymes
that produce quinolinate in plants, and it has often been
stated or implied that there is a strict prokaryotic/eukaryotic
division between quinolinate synthesis pathways (Garavaglia et al., 2003; Bieganowski and Brenner, 2004) and thus
that plants use the tryptophan pathway (Hunt et al., 2004).
Indeed, the tryptophan pathway has been considered an
important route in several plants based on some labelling
studies (reviewed by Arditti and Tarr, 1979). However,
several investigations reported a lack of evidence for
quinolinate synthesis from tryptophan, for example, in
maize and tobacco (Henderson et al., 1958), and other
authors have thus favoured the aspartate pathway while
noting that no enzymes necessary to produce quinolinate
had been found in plants (Mann and Byerrum, 1974b).
Subsequently, it was reported that L-aspartate oxidase
activity could be detected in cotton callus cell extracts
(Hosokawa et al., 1983) and, more recently, analyses of
gene sequences have reported the presence in plant species
of the bacterial pathway of quinolinate synthesis from
aspartate and triose phosphate (Kurnasov et al., 2003;
Katoh and Hashimoto, 2004). Indeed, the Arabidopsis
genome contains, as well as a single gene predicted to
encode QPRT, two sequences that show significant homology to the two bacterial enzymes involved in this pathway (Fig. 1). In contrast, the Arabidopsis genome does not
appear to contain gene sequences that show significant
homology to tryptophan pathway enzymes. Other authors
have noted the presence of genes encoding enzymes of both
the aspartate and the tryptophan pathway in rice (Katoh and
Hashimoto, 2004), but sequences associated with the latter
pathway have been eliminated by subsequent annotation
updates of the genome. Likewise, no sequences of significant homology are found in poplar, Medicago, or Physcomitrella databases. Moreover, a preliminary report has
described the cloning of Arabidopsis homologues for nadA
and nadB, their ability to rescue E. coli mutants deficient in
the respective enzymes, and the presence in plastids of the
encoded proteins (Katoh et al., 2005). These data, together
with the apparent absence of genes encoding enzymes
involved in other possible routes to quinolinate, suggest
that plants obtain this key precursor via the aspartate pathway used by many bacteria (Fig. 1, purple background).
An nadB homologue in plants
L-Aspartate oxidase (EC 1.4.3.16) catalyses the FADdependent oxidation of L-aspartate to iminoaspartate (Fig.
1). FAD is regenerated by either molecular oxygen or
fumarate. A single sequence is annotated L-aspartate
oxidase in Arabidopsis, and homologues are found in
several other plant species. Selected plant and bacterial
protein sequences were used to construct a phylogenetic
tree that reveals a very close relationship between proteins
from angiosperm species, with the moss sequence being
classed slightly apart (Fig. 2A). By contrast, the algal
protein sequence is somewhat intermediate between the
bacterial and plant proteins (Fig. 2A). Bacterial sequences
cluster less tightly than do the plant sequences, probably
because the bacteria for which sequences were analysed
are found in very distinct taxonomic groups.
A sequence alignment was performed to identify conserved amino acid residues in L-aspartate oxidases from
different species (Fig. 2B). The proteins all contain two
large domains, one aspartate oxidase domain in the Nterminal two-thirds of the protein and a succinate dehydrogenase domain in the C-terminal part (Mattevi et al.,
1999). Earlier studies of the cotton enzyme reported the
presence of a permanent cofactor with an apparent molecular mass of 1050 (Hosokawa et al., 1983). This cofactor could be FAD, despite the latter’s smaller molecular
mass, as this nucleotide has been shown to be essential
to the bacterial enzyme. Recent crystallization of the E. coli
enzyme allied with site-directed mutagenesis allowed
identification of amino acid residues involved in the
FAD- and succinate-binding sites (Mattevi et al., 1999;
Bossi et al., 2001; Tedeschi et al., 2001). It is noteworthy
that these amino acids are strictly conserved between plant
and bacterial species (Fig. 2B), consistent with the key
role of the residues in the catalytic properties of the enzyme. Though not shown in Fig. 2B, an analysis of plant
sequences suggests the presence of a chloroplast transit
peptide in L-aspartate oxidase from Arabidopsis, rice,
poplar, and Medicago.
Certain organisms can oxidize aspartate to iminoaspartate using enzymes other than L-aspartate oxidase. In
Thermogata maritima, the nadB gene encodes an NADdependent aspartate dehydrogenase (EC 1.4.1.–), an enzyme with a very different primary protein sequence from
L-aspartate oxidase (Yang et al., 2003). In mammals, a Daspartate oxidase (EC 1.4.3.1) synthesizes iminoaspartate,
and in vitro this enzyme can replace L-aspartate oxidase
in the quinolinate synthase complex (Nasu et al., 1982a).
The physiological role of this enzyme is not in NAD
NAD(P) synthesis and pyridine nucleotide cycling
synthesis but in regulating the concentration of D-aspartate,
a compound present in the brain and some secretory
organs, by converting it to iminoaspartate which then
decomposes to oxaloacetate (Wang et al., 2000; Wolosker
et al., 2000).
An nadA homologue in plants
Iminoaspartate produced by L-aspartate oxidase is combined with dihydroxyacetone phosphate to produce quinolinate, a reaction catalysed by quinolinate synthase (also
called quinolinate synthase ‘A’). Sequences homologous
to the bacterial proteins were found in the Arabidopsis,
rice, poplar, Medicago, and Physcomitrella databases, but
for Chlamydomonas only a partial sequence is available.
For the purposes of sequence alignment, the analysis was
restricted to this part of the protein. Construction of a
phylogenetic tree for the protein sequences generates a clear
distinction between plants and bacteria (Fig. 3A). The five
plant sequences are closely related to each other and also to
the Chlamydomonas sequence. Bacterial sequences form
a cluster in which E. coli and Synechocystis are most
closely related. Plant and bacterial sequences appear more
distinct from each other than for L-aspartate oxidase.
Alignment of plant sequences with the partial Chlamydomonas sequence indicates the presence of several highly
conserved amino acid residues (Fig. 3B). The threedimensional structure of the crystallized quinolinate synthase from Pyrococcus horikoshii shows a triangular
architecture in which conserved amino acids determine
three structurally homologous domains, termed ‘three-fold
repeat of three-layer sandwich’ (Sakuraba et al., 2005).
Comparison of complete bacterial, rice, and Arabidopsis
sequences with the P. horikoshii sequence suggests that
all quinolinate synthases share this unique triangular
architecture, although the plant sequences contain an
additional C-terminal extension.
Recent studies demonstrate that E. coli quinolinate
synthase harbours a [4Fe–4S] cluster (Cicchillo et al.,
2005; Ollagnier de Choudens et al., 2005). The sensitivity
of this cofactor to oxidation probably explains why quinolinate synthase is the site of oxygen poisoning of pyridine
nucleotide synthesis in anaerobic bacteria (Gardner and
Fridovich, 1991; Draczynska-Lusiak and Brown, 1992;
Ollagnier de Choudens et al., 2005). The characteristic
binding motif for [4Fe–4S] clusters is Cys-w-x-Cys-y-zCys, and this is found in the C-terminal region of most
quinolinate synthases from bacteria (Ollagnier de Choudens
et al., 2005), including Synechocystis. No homologous
sequence is found in the plant quinolinate synthase. The
catalytic mechanism and cofactors for the plant enzyme
remain to be identified.
Although less clear for other available plant quinolinate
synthase sequences, the Arabidopsis and poplar enzymes
1609
are predicted to contain an N-terminal plastid targeting
peptide. Together with the chloroplastic localization of
L-aspartate oxidase, this suggests that quinolinate synthesis probably occurs in this organelle. As iminoaspartate is
unstable, L-aspartate oxidase and quinolinate synthase
could occur in close association, although no conclusive
evidence has yet been found in bacteria for the formation of
a complex (Sakuraba et al., 2005).
Compartmentation and transport in
NAD synthesis and metabolism
Total NAD(H) concentrations in leaf cell mitochondria and
cytosol are ;2 and 0.6 mM, respectively, while total
NADP(H) pools are of the order of 0.3 mM in both
compartments (Igamberdiev and Gardeström, 2003).
NAD(H) and NADP(H) pools of isolated chloroplasts
have both been calculated to be ;0.4 mM (Takahama
et al., 1981). Because of their key roles in metabolism in
these compartments and in others such as the peroxisomes,
NAD and NADP show very rapid redox turnover during
processes such as photosynthesis, photorespiration, and
respiration. Numerous small metabolite exchange systems
are able to shuttle redox equivalents rapidly across
chloroplast and mitochondrial membranes and thus link
NAD(P) pools in the different compartments (Raghavendra
and Padmasree, 2003; Scheibe et al., 2005). These involve
triose phosphate/3-phosphoglycerate, malate/oxaloacetate,
and malate/aspartate exchange (Day and Wiskich, 1981;
Journet et al., 1981; Heineke et al., 1991). Glutamate/2oxoglutarate shuttle systems, which are key in nitrogen
assimilation and ammonia reassimilation (Woo et al.,
1987), also potentially involve exchange of redox equivalents. Exchange systems also operate to link peroxisomal
and cytosol NAD pools (Reumann et al., 1994). As well as
indirect redox transfer between the mitochondrial matrix
and the cytosol, evidence has recently been presented that
a cytosolic glycerol-3-phosphate dehydrogenase (G3PDH)
operates with an externally oriented mitochondrial inner
membrane G3PDH:ubiquinone reductase to deliver electrons to the Arabidopsis mitochondrial electron transport
chain, and this system appears to contribute significantly
to cellular redox homeostasis (Shen et al., 2006). Other
externally oriented ‘alternative’ dehydrogenases located
in the inner mitochondrial membrane may also allow
oxidation of NADH or NADPH produced in the cytosol
or intermembrane space (Giegé et al., 2003; Rasmusson
et al., 2004).
In comparison with the redox exchange between compartments, little is known about the transport of NAD
and NADP themselves. As discussed above, available data
suggest that the first three steps of the de novo pathway of
NAD synthesis involve production of quinolinate in the
chloroplast and NaMN formation in the cytosol and
mitochondrion (Fig. 4). In Arabidopsis, only a single gene
1610 Noctor et al.
Fig. 2. Phylogenetic tree and multiple sequence alignment of L-aspartate oxidase (NadB) proteins. (A) The phylogenetic tree was produced from
a multiple sequence alignment using the Clustal W program (http://www.infobiogen.fr/services/analyseq/cgi-bin/clustalw_in.pl). It includes L-aspartate
oxidase protein sequences from Arabidopsis thaliana (AtNadB: GenBank accession number NP_568304), Oryza sativa (OsNadB: XP_464033),
Populus trichocarpa (PtNadB: eugene3.00011954; http://genome.jpgi.pso.org/Potr1/), Synechocystis PCC6803 (SyNadB: NP_442857), E. coli
(EcNadB: NP_754979), B. subtilis (BsNadB: NP_390665), and single homologous sequences extracted from Chlamydomonas reinhardtii
NAD(P) synthesis and pyridine nucleotide cycling
is predicted to encode each of the enzymes necessary for the
subsequent adenylylation and amidation steps. Neither
sequence encodes an apparent transit peptide sequence. In
mammals, three N(a)MNAT isoforms have been characterized: one is found in the nucleus, another outside the nucleus
in an undefined location, and the third in both the cytosol
and mitochondria (Magni et al., 2004). A green fluorescent
protein (GFP)–yeast NAD synthetase fusion construct
showed a diffuse localization in both the cytosol and the
nucleus (Suda et al., 2003). Taken together, available
data suggest that neither plastids nor mitochondria are involved in the last two steps of NAD synthesis, though this
remains to be confirmed (Fig. 4). Based on this view, there
would appear to be a minimum requirement for plastid inner
membrane transporters able to export quinolinate and import
NAD, as well as for mitochondrial import of NAD and
quinolinate for mitochondrial QPRT activity (Sinclair et al.,
2000). Both NAD+ and NADP+ have been shown to cross
plant mitochondrial membranes (Neuburger and Douce,
1983; Bykova and Møller, 2001), but very little genetic and
molecular information is available on NAD transporters.
As they are relatively large, phosphorylated metabolites,
pyridine nucleotides are usually considered to be exclusively intracellular, with virtually no movement between
cells. However, mammalian plasma contains low concentrations of NAD, which are important in signalling through
membrane-bound ADP-ribosyl cyclases such as CD38 that
could act both to produce cADP-R on the extracellular face
of the membrane and to transport it into the cell (Evans
et al., 2005). Recently, a plasma membrane NAD transporter has been shown to operate in a pathogenic intracellular bacterium (Haferkamp et al., 2004). This strain
is unable to synthesize NAD and instead takes it up from
the host cell, probably in exchange for ADP (Haferkamp
et al., 2004). Other bacteria, such as Salmonella typhimurium, are able to take up exogenous NMN via the product
of the pnuC gene, particularly under conditions of low
intracellular NAD+ (Zhu et al., 1991; Penfound and Foster,
1999), where uptake activity could be stimulated by the
regulatory protein NadR (see below). Very recently, two
genes encoding mitochondrial NAD+ transporters have
been identified in yeast where, as in plants, the final steps
of NAD synthesis appear to occur outside the mitochondria
(Todisco et al., 2006). Biochemical characterization of
one of the gene products, NDT1, showed high specificity
for NAD+ (Km = 0.38 mM), with virtually no activity
1611
+
with NADH, NADP , or NADPH, but some activity with
other nucleotides, including NaAD (Todisco et al., 2006).
The protein appears to catalyse unidirectional transport at
low rates and exchange with other nucleotides at faster
rates (Todisco et al., 2006). Genome searches reveal that
homologues of the two yeast mitochondrial transporters
exist in taxonomically diverse organisms, including Arabidopsis, poplar, and rice (Fig. 5). The two Arabidopsis
proteins have 30–31% identity with yeast NDT1 and the
sequences are currently annotated adenine nucleotide
translocator/mitochondrial substrate carrier, with a predicted inner mitochondrial membrane location. It remains
to be seen whether similar proteins exist in the plastid envelope, and whether mitochondrial import of NAD occurs
in exchange for other nucleotides such as NaMN, whose
fate after production by mitochondrial QPRT is unclear
(Fig. 4). Since NAD(H) kinases appear to exist in multiple
compartments (Gallais et al., 2000a; Turner et al., 2004,
2005; Chai et al., 2005), the requirement for NADP
transport is yet to be established.
As well as mitochondrial and chloroplast transporters,
important questions remain concerning nuclear NAD
metabolism and nicotinamide recycling, and their relationship to cytoplasmic processes. Imaging analysis of
NAD(H) suggests that the free nuclear NAD+:NADH ratio
is similar to cytosolic values (Zhang et al., 2002). If
pyridine nucleotides pass readily through nuclear membrane pores, then changes in cellular NAD redox state
or total NAD pools could impact directly on transcription
in animal and yeast cells via redox regulation of the corepressor CtBP (Zhang et al., 2002) or substrate availability for Sir2 and related proteins (Imai et al., 2000; Lin
et al., 2000). Interestingly, the last two enzymes of
NAD synthesis are present in yeast and mammalian nuclei
(Suda et al., 2003; Magni et al., 2004), and significant
resynthesis of NAD could occur within this compartment.
Intercellular transport is another issue. In tobacco,
nicotine is translocated from roots to shoots, and root
synthesis and shoot accumulation are stimulated by stresses
such as wounding (Cane et al., 2005, and references cited
therein). Ricinine is found in the phloem sap of castor bean
(Holfelder et al., 1998), and trigonelline probably moves
between organs in developing mungbean seedlings (Zheng
et al., 2005). Whether any of the other intermediates and
precursors shown in Fig. 1 move significantly between
cells or organs under physiological conditions is unclear.
(CrNadB: http://chlamy.org) and Physcomitrella patens (PpNadB: http://moss.nibb.ac.jp) databases by BLAST search against the other plant and
bacterial sequences shown. A homologous sequence from Medicago truncatula (MtNadB: BAC accession number AC159962, http://www.
medicago.org) was also included in the analysis. Gaps were excluded. The numbers indicate percentage identities between the protein sequences. (B)
Sequence alignment of L-aspartate oxidase proteins. Identical amino acids are highlighted in black and include residues shown for the E. coli enzyme to
be involved in FAD (S22, A25, K44, Q56, G58, A164, and G382) and succinate (G223, E268, R298, H361, and S397) binding, numbered for the Arabidopsis
protein. Sequences in frames indicate similarity. Transit peptides of plant and Chlamydomonas sequences predicted by ChloroP software (http://www.
cbs.dtu.dk/services/ChloroP/) were not taken into account in the alignment.
1612 Noctor et al.
Fig. 3. Phylogenetic tree and multiple sequence alignment of quinolinate synthase (NadA) proteins. (A) The tree was produced as described for
Fig. 2 by comparing quinolinate synthases from A. thaliana (AtNadA: GenBank accession number NP_199832), O. sativa (OsNadA: ABA_97161),
P. trichocarpa (PtNadA: eugene3.00150643; http://genome.jpgi.pso.org/Potr1/), M. truncatula (MtNadA: sequence number TC107403 from the
TIGR database; http://www.tigr.org/tdb/tgi/plant.shtml), P. patens (PpNadA: http://moss.nibb.ac.jp), C. reinhardtii (CrNadA: AAK_54345), Synechocystis PCC6803 (SyNadA: NP_442873), E. coli (EcNadA: NP_752755), and B. subtilis (BsNadA: NP_390663). Gaps were excluded. The
numbers indicate percentage identities between proteins. (B) Sequence alignment of various quinolinate synthase proteins. Identical amino acids are
highlighted in black. Sequences in frames indicate similarity. Because the available Chlamydomonas sequence is incomplete in the C-terminal
domain, only parts of the sequences are aligned.
Stress-induced changes in NAD status
Given the centrality of pyridine nucleotides to cell function,
it seems likely that contents are highly regulated. Operation
of NAD-consuming reactions means that regulation will
be necessary to ensure homeostatic replenishment of NAD
pools. Furthermore, adjustment of NAD pools could play
an important part in environmentally induced ‘dynamic’
signalling, in which the cell is able to sense and respond
to changes in concentrations of NAD and/or precursors
or metabolites (Zhang et al., 2002).
Several NAD-consuming reactions could be important
in stress conditions and/or signalling in interactions with
ROS and other redox components. Most of these reactions
release nicotinamide, which can be recycled to NAD by a
salvage pathway (Fig. 1). One important NAD-consuming
enzyme induced by stress is PARP. This nuclear enzyme
transfers ADP-ribose units from NAD+ to residues on
target proteins, including itself, releasing nicotinamide
and ultimately forming long poly(ADP-ribose) chains that
are important in regulating processes such as DNA repair
(Berglund et al., 1996; Graziani et al., 2005). Hyperactivation of PARP-1 by ROS or nitric oxide has been
shown to deplete NAD pools in mammalian and plant
cells, and decreasing PARP inhibition can diminish
death in both types of cell (Heller et al., 1995; Amor
et al., 1998). Recently, it has been shown that stressinduced PARP activation causes NAD+ and ATP depletion,
and that these effects and the attendant cell death are attenuated in lines in which PARP expression is diminished by
RNA interference (RNAi) (De Block et al., 2005).
As well as producing changes in NAD availability,
adjustments in NAD metabolism could impact on stress
signalling through changes in metabolic intermediates such
as quinolinate or nicotinamide, which are known to be
NAD(P) synthesis and pyridine nucleotide cycling
Fig. 4. Possible subcellular compartmentation of the de novo NAD(P)
synthesis pathway. Established or likely enzymatic conversions are
indicated by solid arrows, uncertain reactions by dashed arrows, and
transport steps by dotted arrows. Other pathways that are not shown could
also be important (see text). Only transport steps that seem to be
necessary based on likely compartmentation of synthetic enzymes are
considered. Other transport reactions could also occur (e.g. transport of
NaAD). Localization of synthetic enzymes is based on available gene
sequences, as well as the reported localization of quinolinate synthesis
enzymes in the chloroplast (Katoh et al., 2005). A tobacco cDNA for
quinolinate phosphoribosyltransferase (QPRT) is predicted to encode
a protein targeted to the mitochondria, although a second transcript
may also be produced from the same gene to produce a shorter protein
(Sinclair et al., 2000). The Arabidopsis gene predicts two proteins, one
lacking the putative mitochondrial target sequence, suggesting that
QPRT is found in both the mitochondria and the cytosol. The subsequent
steps of NAD synthesis are probably located in the cytosol, although other
locations cannot yet be definitively discounted. NADP synthesis occurs
in the chloroplast and the cytosol, and, perhaps, in the mitochondria
(Gallais et al., 2000a; Turner et al., 2004, 2005; Chai et al., 2005).
Mitochondria can import NAD and NADP (Neuburger and Douce,
1983; Bykova and Møller, 2001). For the transporter activities indicated
by ellipses, the only identified protein is the yeast mitochondrial NDT
transporter (Todisco et al., 2006), for which homologous sequences
exist in Arabidopsis, poplar, and rice (see Fig. 5). NDTh, homologue
of the yeast NDT. Other abbreviations as for Fig. 1.
neuroactive compounds in animals (Stone and Addae,
2002; Magni et al., 2004). Quinolinate accumulation in
the brain has been implicated in several neurodegenerative
disorders (Fukuoka et al., 2002, and references therein).
Nicotinamide is known to be an inhibitor of nicotinamidegenerating enzymes such as PARP and Sir2 (Magni et al.,
2004; De Block et al., 2005), and high concentrations have
been shown to influence defence processes in plants
(Berglund et al., 1993, 1996). Besides PARP, other enzymes can modify the biological activity of proteins by
transferring a single ADP-ribose moiety from NAD to
target amino acids. An Arabidopsis mutant, rcd1, which
shows hypersensitive responses to extracellular superoxide, contains a PARP-type NAD-binding signature that
could reflect the mono(ADP-ribose) transferase function
(Ahlfors et al., 2004).
Several literature studies show that NAD pools are
plastic in plants, and can undergo significant changes in
response to the environment. Powdery mildew infection of
barley leaves was reported to be associated with increased
1613
NAD contents, although no evidence was obtained for
increases in synthesis (Ryrie and Scott, 1969). It is evident
from Fig. 1 that NAD contents could be modified by
changes at multiple levels, including degradation and recycling efficiency, as well as de novo synthesis. Within this
context, key points relate to the removal of metabolites
from the recycling pathway for pyridine alkaloid synthesis,
degradation of nicotinamide, or formation of nicotinate
or nicotinamide conjugates (Fig. 1). Enzymes able to
decarboxylate and oxidize both pyridine monocarboxylates
(nicotinate, isonicotinate, and picolinate) and dicarboxylates (quinolinate and dipicolinate) to hydroxylated products are found in bacteria (Uchida et al., 2003). Another
question concerns stress-induced effects on the NAD:
NADP ratio. In cultured tobacco cells expressing a hyperactive calmodulin, increased activity of NAD kinase,
a calmodulin-activated enzyme, was associated with a significant increase in the NADP+:NAD ratio (Harding et al.,
1997). Interestingly, NAD kinase from Bacillus subtilis is
allosterically activated by quinolinate (Garavaglia et al.,
2003), and this could play some role in the co-regulation
of the de novo synthesis of NAD and NADP.
Further evidence pointing to a role for the concentration
of NAD and/or associated metabolites in influencing stress
resistance comes from analysis of tobacco lines deficient
in mitochondrial complex I. These plants respire through
alternative respiratory dehydrogenases. Growth is slowed
but leaf respiration is increased, as is resistance to both
abiotic and biotic stress (Dutilleul et al., 2003a, b).
Enhanced resistance is accompanied by a 2-fold increase
in leaf contents of NAD+ and NADH, without an appreciable increase in NADP+, NADPH, H2O2, ascorbate, or
glutathione (Dutilleul et al., 2003b, 2005). From a physiological point of view, increases in NADH in these plants
can perhaps be rationalized in terms of the relatively
high Km NADH of the alternative matrix-orientated mitochondrial NADH dehydrogenase, which in this line is
likely to be important as a functional replacement of
complex I. The biochemical and molecular processes responsible for increases in NAD in the cytoplasmic male
sterile (CMS) mutant remain unidentified, and further work
is required to determine the extent to which the observed
changes in NAD status in these plants contribute to their
enhanced resistance to stress (Dutilleul et al., 2003b).
The extent to which the redox state of NAD and NADP
pools changes in response to the environment is still
a matter of some debate. Concerning the chloroplast, it is
often assumed that increasing light causes progressive overreduction at the reducing side of photosystem I (PSI),
decreasing the availability of NADP+, the principal oxidant
of ferredoxin, and so favouring diversion of electrons to
oxygen to produce ROS. However, the stromal redox state
is rather constant in response to changes in irradiance,
though less so when metabolism strongly limits electron
transport, for example, at low temperatures or if both CO2
1614 Noctor et al.
Fig. 5. Phylogenetic tree and multiple sequence alignment of yeast mitochondrial NAD+ transporters NDT1 and NDT2 (Todisco et al., 2006) and
homologous protein sequences in other organisms. (A) The tree was produced as described for Figs 2 and 3, by BLAST search with Saccharomyces
cerevisiae NDT1 as probe. (B) Sequence alignment of yeast NDTs and homologous sequences in other organisms. Identical amino acids are highlighted
in black. Sequences in frames indicate similarity. Key to sequences: ScNDT1, Saccharomyces cerevisiae NP_012260; ScNDT2, S. cerevisiae
NAD(P) synthesis and pyridine nucleotide cycling
and O2 are artificially decreased (Harbinson et al., 1990).
Direct measurements of fractionated pea leaf protoplasts
showed that cytosolic NADP redox states were relatively
independent of light, dark, or CO2 availability (Igamberdiev
and Gardeström, 2003). However, cytosolic NAD and
both mitochondrial NAD and NADP pools were more
reduced in the light, particularly under conditions favouring
photorespiration (Igamberdiev and Gardeström, 2003). It
was proposed that these changes are an important factor in
mediating light-induced redirection of leaf metabolism,
particularly via inhibition of the mitochondrial NAD-ICDH
caused by more reduced NAD pools (Igamberdiev and
Gardeström, 2003). Such phenomena could be at least
partly responsible for some of the changes in metabolite
profiles observed in the tobacco CMS mutant, where
increases in total leaf NAD+ and NADH are associated
with enhanced nitrate reduction and an increased citrate:2oxoglutarate ratio despite enhanced extractable NADICDH activity (Dutilleul et al., 2005). However, as for
almost all other similar work, pyridine nucleotide measurements in these studies represent total nucleotide pools, i.e.
they include nucleotides that are bound to proteins as well
as the more biochemically relevant ‘free’ pools. Recent
analyses of isolated potato tuber mitochondria in short-term
experiments have led to the conclusion that free mitochondrial NADH is maintained at a more constant value than
total NAD+ and NAD, at least in organelles isolated from
non-photosynthetic cells (Kasimova et al., 2006). Several
other recent studies using leaf systems indicate that changes
in cytosolic and mitochondrial NAD status are important
in the light–dark regulation of nitrate reduction and respiratory pathways associated with nitrogen assimilation
(Kaiser et al., 2002; Igamberdiev and Gardeström, 2003;
Rachmilevitch et al., 2004; Dutilleul et al., 2005).
Notwithstanding the attention paid to chloroplastic
events using non-invasive techniques such as chlorophyll
fluorescence, the impact of stress on NAD(P) pools and
redox states in the different compartments remains unresolved. NAD(P)H oxidases are among the key players in
ROS signalling, and PARP (and perhaps other NADconsuming enzymes) can be activated by stress. In this
context, key questions relate, first, to possible links between NAD degradation/(re)synthesis and redox states of
NAD(P) and, secondly, to what extent NAD(P) pools in
different compartments are autonomous or interdependent. Given that NAD(P)H oxidases use cytosolic reductant and several NAD-consuming enzymes are located
1615
in the nucleus, the cytosol and nucleus could be the key
locations for NAD-linked signalling. Further information
is required to establish whether chloroplast and mitochondrial pools might be relatively insulated from potentially
larger fluctuations in cytosolic NAD. Within this context,
the location of the different reactions of the pyridine cycling
pathway is an important consideration.
Regulation of NAD contents
What are the mechanisms that could act to regulate NAD
synthesis in plants? In bacteria, in which NAD metabolism
involves similar enzyme activities to those found in plants,
regulation occurs at transcriptional and post-translational
levels. The bacterial nadA, nadB, and nadC genes are
grouped in the nad operon under the control of a single
promoter (Penfound and Foster, 1996). In S. typhimurium,
transcription of nadA, nadB, and pncB (encoding NaPRT)
is repressed by the DNA-binding function of the repressor protein NadR (Penfound and Foster, 1999). Binding of
NadR to the nadA operator is highly dependent on NAD+,
and uninfluenced by physiological concentrations of quinolinate, nicotinate, nicotinamide, NaMN, NMN, NaAD,
NADH, or NADP+ (Penfound and Foster, 1999). NadR is an
intriguing protein for which two additional activities have
been described. First, as well as the DNA-binding repressor
function that is favoured in NAD+-replete cells, NadR could
act in NAD+-deficient cells as a transport facilitator in
conjunction with the pnuC gene product, encoding a NMN
transporter protein (Penfound and Foster, 1999). Secondly,
in addition to repressor and transport functions, NadR also
has dual nicotinamide riboside kinase/NMNAT activity.
These activities have been described for NadR from several
bacteria, but seem to be particularly important in Haemophilus influenzae. In this species, which lacks the nadA,
nadB, nadC, and pncB genes and thus many of the enzymes
shown in Fig. 1, the DNA-binding domain is absent
from NadR, but the presence of the enzymatic domains
enables NAD+ to be produced from absorbed nicotinamide
riboside (Singh et al., 2002). In B. subtilis, no homologues
of NadR are found, but another repressor, YrxA, regulates
nadA and nadB transcription (Rossolillo et al., 2005). In
this bacterium, however, the quinolinate synthesis pathway is regulated antagonistically with the pyridine
cycling pathway because the metabolite that enhances
the DNA-binding activity of YrxA is nicotinate, not
NAD, and a further difference from the NadR system is
NP_010910; AtA, ScNDT1 homologue A from A. thaliana, protein NP_564233, gene At1g25380; AtB, ScNDT1 homologue B from A. thaliana,
protein NP_566102, gene At2g47490; PtA, ScNDT1 homologue A from P. trichocarpa, fgenesh1_pm.C_LG_X000398; PtB, ScNDT1 homologue B
from P. trichocarpa, eugene3.00140704; OsA, ScNDT1 homologue A from O. sativa, protein BAD73272 (currently annotated as a folate transporter,
gene identifier OS01G32980); OsB, Sc NDT1 homologue B from O. sativa, protein AAV43843, gene OS05G28870 (currently annotated as
a mitochondrial carrier protein); Hs, Homo sapiens protein NP_110407 (currently annotated as a folate transporter); Xl, Xenopus laevis protein
AAH87370; DrA, ScNDT1 homologue A from Danio rerio (Zebrafish), protein AAH90770; DrB, ScNDT1 homologue B from D. rerio, protein
NP_956550; and Dm, ScNDT1 homologue from Drosophila melanogaster, protein AAM68821.
1616 Noctor et al.
that YrxA does not repress pncB transcription (Rossolillo
et al., 2005). The end-result is that quinolinate synthesis is
only activated in B. subtilis when nicotinate is not available. Analysis of available sequences suggests that no
NadR or YrxA homologues are to be found in plants, and
the importance of transcriptional control of NAD synthesis
is unclear.
In E. coli, NAD+ and NADH concentrations can posttranslationally regulate NAD synthesis by feedback inhibition. An in vitro assay of the E. coli nadA and nadB
gene products in combination showed that NAD concentrations close to physiological (1–3 mM) strongly inhibited
quinolinate formation, while NADP had much less effect
(Griffith et al., 1975). It was subsequently shown that the
first enzyme, L-aspartate oxidase, is inhibited by NAD+
(competitively with the prosthetic group FAD), while the
second enzyme, quinolinate synthase, is inhibited by both
NAD+ and NADH (Nasu et al., 1982b). Thus, decreases
in NAD+ should release feedback inhibition over quinolinate synthesis. If the plant homologues are subject to
similar regulation, a key point will concern the relationship
between NAD status outside and within the chloroplast,
where quinolinate synthesis is probably located (Fig. 4). It
is possible that the most important adjustments of quinolinate synthesis could occur through changes in protein
concentrations, mediated by transcriptional or posttranscriptional processes that result from changes in cytosolic or nuclear NAD concentration, NAD+:NADH ratios,
or other signal. Quinolinate or nicotinate could also have
some direct inhibitory effect on the enzyme activities.
Post-translational mechanisms acting through covalent
modification of enzymes or through protein–protein interactions are also likely to be important in regulating the web
of reactions shown in Fig. 1. Certain plant NAD kinases are
regulated by calmodulin (Harding et al., 1997) and also
perhaps by thiol–disulphide exchange (Delumeau et al.,
2000). An important issue is the extent to which quinolinate
synthesis in plants requires complex formation between
L-aspartate oxidase and quinolinate synthase. Reversible
complex formation between enzymes is known to be a key
mode of regulation of plant metabolic pathways such as
cysteine synthesis from sulphide and serine (Droux, 2004).
An important question concerns the importance of
quinolinate synthesis compared with nicotinamide recycling in different plant tissues or environmental conditions,
or at specific stages of development. Labelling studies
suggest that NAD and NADP pools turn over within a few
hours even under optimal conditions (Ryrie and Scott,
1969). Turnover times could be significantly shortened by
increased NAD utilization, and in these conditions the
pyridine recycling path may be particularly important.
NaPRT is considered to be the limiting enzyme in the
recycling pathway in bacteria, and overexpression of this
enzyme in E. coli can cause a 5-fold increase in cellular
NAD concentrations in the presence of added nicotinate
(Wubbolts et al., 1990). More recently, it was shown that
expression of the S. typhimurium enzyme in E. coli
increased NAD pools ;1.5-fold and that this was associated with increased NAD+:NADH ratios (Berrı́os-Rivera
et al., 2002). In barley leaf tissue, NAD was labelled at
similar rates from [14C]nicotinate and [14C]quinolinate
(Ryrie and Scott, 1969). It might be predicted that rapidly
growing tissue would require an active de novo synthesis
from quinolinate whereas the recycling pathway could
perhaps be more important in mature tissues such as fully
expanded leaves. In theory, the de novo synthesis pathway
could be dispensable in expanded cells if recycling is close
to 100% efficient. One obvious factor that would affect
recycling efficiency is the extent of pyridine ‘leakage’ from
the pathway, for instance via pyridine ring degradation.
A second factor impacting on pyridine cycling is the use
of pyridine rings to produce compounds other than
nucleotides. Regulation in such systems probably differs
from that operating in cells in which the primary pyridine
compounds are NAD and NADP. Tissue NAD and NADP
pools in plants are relatively small, typically of the order
of 20–100 nmol gÿ1 fresh weight (Dutilleul et al., 2005;
Shen et al., 2006). By contrast, pyridine alkaloids can accumulate much more strongly in certain species (Mann
and Byerrum, 1974b). Trigonelline contents in coffee
leaves can reach 20 lmol gÿ1 fresh weight (Zheng et al.,
2004). Tobacco presents an example of differential regulation of the NAD synthesis pathway between tissues that
produce nicotine and those that do not. Analysis of QPRT
in Nicotiana species that predominantly produce nicotine
(N. tabacum and N. sylvestris) showed that transcripts were
confined to roots (where nicotine is synthesized) while
QPRT was expressed in both roots and leaves in N. glauca,
in which the principal pyridine alkaloid is anabasine
(Sinclair et al., 2000). Wounding of aerial tissues greatly
increased QPRT transcripts in all three species, although
it is notable that induction was confined to the roots in
the nicotine-producing species (Sinclair et al., 2000). Preliminary data on L-aspartate oxidase and quinolinate
synthase also appear to suggest that, in tobacco, the de
novo pathway of NaMN synthesis is most highly expressed
in conditions in which nicotine synthesis is stimulated. Like
QPRT, these enzymes appear to be most expressed in the
tobacco root and are similarly induced by methyl jasmonate
(Katoh and Hashimoto, 2004). In contrast, they are not
induced by jasmonate treatment of tobacco leaves (Katoh
and Hashimoto, 2004). In castor bean seedlings accumulating ricinine, extractable QPRT activities were much
higher than those of NaPRT, but in mature leaves, in which
ricinine synthesis is much less significant, the two activities
were similar (Mann and Byerrum, 1974b). The same study
reported that of the two enzymes in tobacco, QPRT was
the more active, whereas in other species the two activities were comparable (Mann and Byerrum, 1974b).
High or increased activity of quinolinate production and
NAD(P) synthesis and pyridine nucleotide cycling
use presumably reflects enhanced production of alkaloids
as feeding deterrents. Whether the de novo pathway of
NAD synthesis has any role to play in more general
conditions of stress, in which NAD metabolism is stimulated or perturbed, remains to be seen.
Conclusions and perspectives
Work in animal and yeast systems on the signalling roles of
NAD and derivatives has rekindled interest in pyridine
nucleotide synthesis and turnover. Recent studies in plants
suggest that NAD contents are both flexible and potentially
important in determining cell fate. These developments
are introducing a new dimension into studies of pyridine
nucleotides, in which studies of synthesis and turnover
are examined alongside the hitherto dominant analysis of
redox state and redox flux. In relation to stress physiology,
a key outstanding question is the extent to which plants
adjust their NAD pools in different conditions. Linked to
this issue is the eludication of the roles of enzymes such
as PARPs, SIRTUINS, and cADP-R in plants, and the
impact of these activities on NAD contents, turnover, and
synthesis. This review has focused specifically on the production and metabolism of the pyridine ring of NAD(P),
but it is clear that future work on NAD will also have
to consider co-ordination of purine and pyridine nucleotide metabolism. As well as representing the major product
of NAD(P)H oxidation in biological energy transduction,
ATP is also required to produce the adenosine monophosphate moiety of NAD(P) and as a phosphate donor in
several reactions shown in Fig. 1.
Together, molecular, genetic and biochemical studies are
beginning to elucidate the web of reactions underpinning
NAD synthesis and pyridine recycling in plants. Current
knowledge strongly suggests that like bacteria, plants make
quinolinate from aspartate and triose phosphate. Genes
enabling quinolinate synthesis from tryptophan appear to
be absent from plant and Chlamydomonas genomes.
Certain pathogenic bacteria have been identified that use
the tryptophan pathway (Kurnasov et al., 2003), but no
organism has been unambiguously identified that uses
both routes for quinolinate synthesis. Taken together, the
genomic data suggest that there is no simple schism between prokaryotic and eukaryotic organisms with respect
to quinolinate synthesis. It is intriguing that enzymes
catalysing the two steps seem to be each encoded by
a unique gene and that both sequences are relatively highly
conserved between divergent taxonomic groups. This
suggests a strong selection pressure on this pathway, reflecting its fundamental importance in plant and bacterial
cell life. Indeed, the pathway has been proposed as an
example of how abiotic chemical evolution may have
occurred in pre-cellular systems (Wächtershäuser, 1988).
One element of this engaging theory is chemical autocatalysis through mechanisms that are independent of
1617
polypeptide enzymes, and it is suggested that quinolinate
formation could have been autocatalytically stimulated
by NaMN-dependent oxidation of aspartate to iminoaspartate (Wächtershäuser, 1988). Following the evolution of
enzyme catalysts, the flavoprotein L-aspartate oxidase
(or precursor) would have taken over this role.
The plant pyridine cycling pathway seems to have more
in common with the bacterial pathway than with animal and
fungal models. Additional reactions could also be present
in plants, enabling enzymes to be by-passed under certain
conditions and hence conferring potentially important
metabolic flexibility. Reverse genetics approaches will be
necessary to establish the necessity or otherwise of such
reactions. The question of enzyme specificity is also key.
Biochemical characterization of the purified enzymes will
be required to confirm the putative preferences of the
different phosphoribosyltransferases and adenylyltransferases. It remains to be seen whether the NaMN adenylyltransferase (a single annotated gene in Arabidopsis)
has any role in direct production of NAD from NMN.
Metabolite profiling could also provide useful complementary information on the presence of given reactions
in plant tissues, and understanding the integration of
NAD(P) synthesis and turnover will require confirmation
of the subcellular localization of enzymes and characterization of intracellular transport systems.
Acknowledgements
We thank Vincent Thareau for expert advice on gene sequence
analysis, and an anonymous reviewer for constructive comments that
helped to improve the manuscript in revision.
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