The FEBS Journal

Volume 274, Issue 1 p. 1-22

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Vitamin C

Biosynthesis, recycling and degradation in mammals

Carole L. Linster,

Carole L. Linster

Université Catholique de Louvain, Christian de Duve Institute of Cellular Pathology, Brussels, Belgium

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Emile Van Schaftingen,

Emile Van Schaftingen

Université Catholique de Louvain, Christian de Duve Institute of Cellular Pathology, Brussels, Belgium

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Carole L. Linster,

Carole L. Linster

Université Catholique de Louvain, Christian de Duve Institute of Cellular Pathology, Brussels, Belgium

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Emile Van Schaftingen,

Emile Van Schaftingen

Université Catholique de Louvain, Christian de Duve Institute of Cellular Pathology, Brussels, Belgium

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First published: 06 December 2006

https://doi.org/10.1111/j.1742-4658.2006.05607.x

Citations: 556

E. Van Schaftingen, Laboratory of Physiological Chemistry, UCL-ICP, Avenue Hippocrate 75, B-1200 Brussels, Belgium
Fax: +32 27647598
Tel: +32 27647564
E-mail: [email protected]

C. L. Linster, The Department of Chemistry and Biochemistry and the Molecular Biology Institute, University of California, Los Angeles, CA 90095-1569, USA
Fax: +1 310 825 1968
Tel: +1 310 825 3137
E-mail: [email protected]

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Abstract

Vitamin C, a reducing agent and antioxidant, is a cofactor in reactions catalyzed by Cu⁺-dependent monooxygenases and Fe²⁺-dependent dioxygenases. It is synthesized, in vertebrates having this capacity, from d-glucuronate. The latter is formed through direct hydrolysis of uridine diphosphate (UDP)-glucuronate by enzyme(s) bound to the endoplasmic reticulum membrane, sharing many properties with, and most likely identical to, UDP-glucuronosyltransferases. Non-glucuronidable xenobiotics (aminopyrine, metyrapone, chloretone and others) stimulate the enzymatic hydrolysis of UDP-glucuronate, accounting for their effect to increase vitamin C formation in vivo. Glucuronate is converted to l-gulonate by aldehyde reductase, an enzyme of the aldo-keto reductase superfamily. l-Gulonate is converted to l-gulonolactone by a lactonase identified as SMP30 or regucalcin, whose absence in mice leads to vitamin C deficiency. The last step in the pathway of vitamin C synthesis is the oxidation of l-gulonolactone to l-ascorbic acid by l-gulonolactone oxidase, an enzyme associated with the endoplasmic reticulum membrane and deficient in man, guinea pig and other species due to mutations in its gene. Another fate of glucuronate is its conversion to d-xylulose in a five-step pathway, the pentose pathway, involving identified oxidoreductases and an unknown decarboxylase. Semidehydroascorbate, a major oxidation product of vitamin C, is reconverted to ascorbate in the cytosol by cytochrome b₅ reductase and thioredoxin reductase in reactions involving NADH and NADPH, respectively. Transmembrane electron transfer systems using ascorbate or NADH as electron donors serve to reduce semidehydroascorbate present in neuroendocrine secretory vesicles and in the extracellular medium. Dehydroascorbate, the fully oxidized form of vitamin C, is reduced spontaneously by glutathione, as well as enzymatically in reactions using glutathione or NADPH. The degradation of vitamin C in mammals is initiated by the hydrolysis of dehydroascorbate to 2,3-diketo-l-gulonate, which is spontaneously degraded to oxalate, CO₂ and l-erythrulose. This is at variance with bacteria such as Escherichia coli, which have enzymatic degradation pathways for ascorbate and probably also dehydroascorbate.

Abbreviations

BSO: l-buthionine-(S,R)-sulfoximine
DHA: dehydroascorbate
2,3-DKG: 2,3-diketo-l-gulonate
FAD: flavin adenine dinucleotide
GLO: l-gulonolactone oxidase
GSH: glutathione (reduced form)
GST: glutathione S-transferase
GSTO: Omega class glutathione S-transferase
PDI: protein disulfide isomerase
SDA: semidehydroascorbate
UDP: uridine diphosphate
UGT: UDP-glucuronosyltransferase

Vitamin C (or l-ascorbic acid; hereafter, ‘ascorbic acid’ and ‘ascorbate’ will always refer to ‘l-ascorbic acid’ and ‘l-ascorbate’) is unique among vitamins for several reasons. It is present in various foods, particularly of plant origin, in quantities (typically 10–100 mg/100 g [1]) that are several orders of magnitude higher than those of other vitamins. This is certainly related to the facts that it is formed from sugars, which are abundant compounds, and that its biochemical synthesis is rather simple. Another unique aspect of ascorbic acid is that it is a vitamin for only a few vertebrate species, those which have lost the capacity to synthesize it. From a structural point of view, it is also one of the rare compounds containing a hydroxyl group that is so acidic as to be completely dissociated at neutral pH (carbon-3 hydroxyl pK_a = 4.2). This is related to the fact that ascorbic acid comprises two conjugated double bonds and that a resonance form can be written for the deprotonated monoanionic form (Fig. 1). Resonance forms can also be written for the form of vitamin C that has lost one electron (Fig. 1), making the radical semidehydroascorbate (SDA) much more stable, and thus much less reactive, than most other free radicals [2]. Vitamin C is therefore able to play the role of a free-radical scavenger [3], reacting with highly ‘aggressive’ (oxidizing) species to replace them by a much less reactive and, moreover, enzymatically recyclable one, SDA. Ascorbate is certainly the most abundant water-soluble compound acting in one-electron reactions, and this is most probably why it plays the role of a cofactor in reactions catalyzed by a number of metal-dependent oxygenases. The Cu⁺-dependent monooxygenases peptidylglycine α-amidating monooxygenase and dopamine β-hydroxylase convert two ascorbate molecules to two SDAs per catalytic cycle [4]. In the case of Fe²⁺/α-ketoglutarate-dependent dioxygenases (e.g. collagen prolyl and lysyl hydroxylases, the two hydroxylases involved in carnitine biosynthesis [5], the asparaginyl hydroxylase that modifies hypoxia-inducible factor 1 (HIF-1) [6]), ascorbate most probably serves to reconvert inactive, Fe³⁺-containing enzyme (which results from abortive catalytic cycles) to the active, Fe²⁺-containing form [5]. Because of these important roles, it is not surprising that vitamin C deficiency leads to a debilitating disorder, scurvy, in man and in animals unable to synthesize the vitamin.

Details are in the caption following the image — **Figure 1**
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The three redox states of vitamin C (ascorbate, fully reduced form; SDA, monooxidized form; DHA, fully oxidized form), and stabilization of the ascorbate monoanion and SDA by electron delocalization. SDA, semidehydroascorbate; DHA, dehydroascorbate.

Important progress has been made recently in our understanding of the synthesis and the recycling of vitamin C, and a novel pathway has been described for the degradation of vitamin C in bacteria. This forms the subject of this review. Vitamin C transport, which has also witnessed important developments lately, is only briefly alluded to in the following paragraph, as other recent reviews are available [7–9].

Ascorbate entry into mammalian cells is energy-dependent, being effected by two distinct Na⁺-dependent cotransporters, SVCT1 and SVCT2, which show distinct tissue distributions. Interestingly, targeted deletion of the widely distributed SVCT2 is lethal in mice [10], further underlining the importance of vitamin C. Dehydroascorbate (DHA; see Fig. 1) is transported by glucose transporters, particularly GLUT1, GLUT3 and GLUT4 [9], and is therefore not energetically driven. However, intracellular DHA is readily converted to ascorbate (see ‘Recycling of vitamin C’) and this highly favourable reductive step drives DHA uptake by the cell. There are also mechanisms allowing the efflux of ascorbate from cells [7], e.g. from enterocytes during intestinal absorption and from liver cells, which in many mammals produce ascorbate, but the molecular identity of the proteins involved in this process is not yet firmly established.

Formation of vitamin C in mammals and other vertebrates

Outline of the pathway

Ascorbate is synthesized by many vertebrates. The occurrence of ascorbate biosynthesis in sea lamprey [11] suggests that this trait appeared early in the evolutionary history of fishes (590–500 million years ago), i.e. prior to terrestrial vertebrate emergence [12]. The biosynthetic capacity has, however, subsequently been lost in a number of species, such as teleost fishes, passeriform birds, bats (intriguingly, not only the fruit-eating ones, but also others, feeding on blood or insects [13]), guinea pigs, and primates including humans, for whom ascorbate has thus become a vitamin. Fish, amphibians and reptiles synthesize ascorbate in the kidney, whereas mammals produce it in the liver [11,14].

Vitamin C is also formed by all plant species studied so far [15] and yeasts produce d-erythroascorbate, a C₅ analogue of ascorbate [16]. Interestingly, very different pathways have evolved for vitamin C biosynthesis in animals, plants and fungi. In animals, d-glucuronate, derived from UDP-glucuronate, is reduced to l-gulonate, which leads to inversion of the numbering of the carbon chain (‘inversion of configuration’) since the aldehyde function of d-glucuronate (C1) becomes a hydroxymethyl group in the resulting l-gulonate (Fig. 2; see [16] for a review of the early literature). The latter is converted to its lactone, which is oxidized to l-ascorbate by l-gulonolactone oxidase (GLO). In plants, the pathway starts with GDP-d-mannose, which is converted (without change in carbon numbering) to l-galactonolactone, the substrate for the plant homologue of GLO, l-galactonolactone dehydrogenase [15]. The synthesis of d-erythroascorbate in yeasts proceeds from d-arabinose [16], but the mechanisms of formation of the latter have not been elucidated.

Effect of xenobiotics on vitamin C formation

The regulation of vitamin C formation by xenobiotics is described here, because it helps to understand the mechanism of d-glucuronate formation. Other aspects of this regulation are described in a separate section.

It was already observed in the 1940s that administration of a series of xenobiotics to animals was followed by enhanced urinary excretion of ascorbate. The stimulatory effect is shared by a wide variety of structurally unrelated substances such as barbiturates, paraldehyde, chloretone, aminopyrine, antipyrine, 3-methylcholanthrene, polychlorinated biphenyls (PCB) and 1,1,1-trichloro-2,2-bis(p-chlorophenyl)ethane (DDT) [17–19]. Turnover rate studies using radiolabelled ascorbate indicated that the amount of ascorbate synthesized per day was four- to eight-fold higher in chloretone- or pentobarbital-treated rats than in untreated animals [20]. Furthermore, chloretone and barbital were shown to greatly stimulate the incorporation of radiolabelled glucose into urinary glucuronate and ascorbate [21]. As barbital was found to be neither metabolized nor conjugated, but excreted unchanged in urine, its stimulatory effect on urinary glucuronate and ascorbate excretion was proposed to be unrelated to any detoxification mechanism. This view was further supported by the observation that compounds such as borneol, α-naphthol and phenolphthalein, known to be primarily excreted as glucuronides, had essentially no effect on ascorbate excretion [21]. Furthermore, the findings that the in vivo conversion of both d-glucose and d-galactose to glucuronate and ascorbate was increased by xenobiotics [22], but that this was not the case for the conversion of radiolabelled d-glucuronolactone or l-gulonolactone to ascorbate [23], suggested that stimulation occurs at a step between UDP-glucose and d-glucuronolactone in the ascorbate biosynthesis pathway.

Many of the following investigations on this subject studied the effect of agents stimulating vitamin C formation on the activity levels of several enzymes potentially implicated in ascorbate synthesis. UDP-glucose dehydrogenase and UDP-glucuronosyltransferases (UGTs) were found to be induced by some agents, although not by all of them [18,24]. A study using Gunn rats [25] provided highly suggestive evidence for the involvement of UGTs in the formation of vitamin C. Gunn rats are deficient in UGT isoforms of the UGT1A family, but not of the UGT2 family [26,27]. 3-Methylcholanthrene, an inducer of UGTs of the UGT1A family, increased urinary excretion of ascorbate in normal rats (five-fold) and heterozygous Gunn rats (two-fold), but not in homozygous Gunn rats. However, treatment with phenobarbital (an inducer of isoforms of the UGT2 family) increased the urinary excretion of ascorbate in normal and homozygous Gunn rats. Taken together, these results indicate that UGT isoforms of the UGT1A, but probably also of the UGT2 family are involved in ascorbate biosynthesis, possibly by forming a glucuronidated intermediate that would be hydrolyzed by microsomal β-glucuronidase or, as suggested below, by catalyzing the hydrolysis of UDP-glucuronate to UDP and glucuronate (mechanisms b and c described in the next subsection).

These effects on enzyme levels require increased gene transcription and new protein synthesis, which implies that the effect of xenobiotics on vitamin C formation is a long-term effect. However, recent work on isolated hepatocytes demonstrated that the effect of xenobiotics (e.g. aminopyrine, metyrapone, chloretone) on the formation of vitamin C and of its precursor, free glucuronate, occurs in a matter of minutes [28]. The increase in free glucuronate formation, which was best observed in the presence of an inhibitor of the downstream enzyme, glucuronate reductase, was already apparent after 5 min and reached up to 15-fold. It was accompanied by a decrease in the UDP-glucuronate level, but little if any change in the concentration of UDP-glucose, indicating that the effect of xenobiotics on vitamin C formation consists in a short-term effect involving an increase of the conversion of UDP-glucuronate to glucuronate (Fig. 2) and not an increase in the concentration of upstream precursors of glucuronate (‘push-effect’). Most of the stimulating agents did not give rise to detectable amounts of β-glucuronides, arguing against the involvement of a glucuronidation-deglucuronidation cycle in the stimulation of ascorbate formation (see below). It may be interesting to notice that up to ∼100 nmol hexose units·min⁻¹·g⁻¹ liver are channelled towards glucuronate formation in the presence of saturating concentrations of stimulating xenobiotics [28].

Formation of glucuronate from UDP-glucuronate

The formation of glucuronate from UDP-glucuronate could hypothetically involve (a) the cleavage of UDP-glucuronate to glucuronate 1-phosphate, followed by dephosphorylation of the latter by a glucuronate-1-phosphatase [29,30]; (b) the formation of a glucuronidated intermediate, on an exogenous or endogenous acceptor, followed by its hydrolysis by β-glucuronidase [31] or esterases (which could hydrolyze acyl-glucuronides); or (c) direct hydrolysis of UDP-glucuronate to UDP and glucuronate (Fig. 2). As explained below, recent work performed on liver microsomes supports the third mechanism.

As a follow-up of the work showing that a series of nonglucuronidable xenobiotics rapidly stimulate the formation of glucuronate in isolated hepatocytes [28], it was found that the same xenobiotics also stimulated the formation of glucuronate from UDP-glucuronate in liver cell-free extracts enriched with ATP or in liver microsomes supplemented with ATP and a heat-stable cofactor identified as coenzyme A [32]. Quantitatively, the formation of glucuronate observed under these conditions accounted for the formation of glucuronate observed in intact cells, indicating that glucuronate is formed from UDP-glucuronate by a microsomal enzyme. This enzyme is most probably present in the endoplasmic reticulum as, similarly to UGTs, it is stimulated by UDP-N-acetylglucosamine, which enhances the transport of UDP-glucuronate into vesicles derived from the endoplasmic reticulum [33].

Although rat liver microsomal preparations hydrolyze UDP-glucuronate to glucuronate 1-phosphate (presumably because they are contaminated with plasma membrane fragments, which contain a highly active nucleotide pyrophosphatase [34,35]), their glucuronate 1-phosphate phosphatase activity is insufficient to account for the formation of free glucuronate by this preparation [32]. Furthermore, the formation and hydrolysis of glucuronate 1-phosphate are unaffected by the nonglucuronidable xenobiotics under conditions under which glucuronate formation is stimulated approximately three-fold. These and other arguments [32] exclude mechanism (a).

Mechanism (b), which is supported by the observations made on Gunn rats [25], is ruled out by the fact that glucuronate formation from UDP-glucuronate occurs in rat liver microsomes in the absence of UGT substrates and is actually inhibited by such substrates (see below). Furthermore, inhibitors of β-glucuronidase and esterases do not affect the formation of glucuronate from UDP-glucuronate by microsomal preparations [32], ruling out also the involvement of an endogenous glucuronide acceptor that would still be present in washed liver microsomes.

Taken together, these observations lead to the conclusion that glucuronate is formed by direct hydrolysis of UDP-glucuronate by a UDP-glucuronidase (mechanism c). The findings that UDP-glucuronidase is similarly sensitive to various detergents as UGTs and that it is inhibited by UGT substrates suggest that it is a side-activity of these transferases (representing ∼5% of the transferase activity) [32]. The identification of UGTs as the UDP-glucuronidase also allows one to reconcile mechanism (c) with the observations made on Gunn rats [25]. In the only study that focuses on the UDP-glucuronate hydrolase activity of a purified UGT, Hochman and Zakim [36] found that GT_2P had a minor UDP-glucuronidase activity, which could be stimulated by phenylethers and lysophosphatidylcholines up to ∼0.03% of the transferase activity. When transfected into human embryonic kidney cells, human UGT1A6 displayed a hydrolase to transferase activity ratio of 0.4% under certain conditions [32], which is still about one order of magnitude lower than the ratio observed in liver microsomes. It is likely that the ability of UGTs to hydrolyze UDP-glucuronate varies among isoforms and depends on the phospholipidic environment. This last point may explain the observation that UDP-glucuronidase is inhibited in rat liver microsomes by the addition of ATP and coenzyme A [32], as the latter combination of cofactors could allow the reesterification of lipids in a subcellular fraction known to contain free fatty acids, acyl-CoA synthetase and acyltransferases. Further work is obviously needed to establish which UGT isoforms are involved in the formation of free glucuronate and the conditions under which they are able to do so.

Glucuronate reductase (aldehyde reductase)

The reduction of d-glucuronate to l-gulonate is catalyzed by an NADPH-dependent reductase, with broad specificity, known as aldehyde reductase or TPN l-hexonate dehydrogenase in the older literature [37] and now referred to as aldo-keto reductase 1A1 (AKR1A1) for the human enzyme [38]. K_m values of 0.33 and 0.69 mm were obtained for d-glucuronate and d-glucuronolactone, respectively, and these compounds are converted by aldehyde reductase to l-gulonate and l-gulonolactone, respectively [37].

Aldehyde reductase belongs to the large group of monomeric NADPH-dependent oxidoreductases, known as aldo-keto reductases, which comprise many members in the human genome, including aldose reductases (the closest homologues of human aldehyde reductase, sharing ∼65% sequence identity [39]) and hydroxysteroid dehydrogenases [38]. These enzymes display broad substrate specificities and it would therefore not be surprising that, besides aldehyde reductase, other members of the aldo-keto reductase superfamily participate in the reduction of d-glucuronate. Aldose reductase appears to be much less efficient than aldehyde reductase in this respect [40]. Furthermore, as it is barely expressed in liver [41], it is unlikely that it contributes significantly to vitamin C formation. Aldose reductase inhibitors, which have been developed in the hope of preventing diabetic complications by blocking the enhanced formation of sorbitol from glucose in hyperglycaemic states, usually cross-react with aldehyde reductase [42,43]. Thus, sorbinil, an inhibitor of both aldehyde reductase and aldose reductase [42,43], was shown to block the conversion of glucuronate to downstream metabolites (Fig. 2) and inhibit the formation of vitamin C in isolated rat hepatocytes [28], supporting the involvement of aldehyde reductase in ascorbate synthesis.

Urono- and gulonolactonase

As discussed in the next paragraph, the enzyme that forms vitamin C acts on the lactone form of l-gulonate. The conversion of d-glucuronate to l-gulonolactone requires the action of two enzymes, a reductase and a lactonase, proceeding either via d-glucuronolactone if the lactonization is the first step, or via l-gulonate if the first reaction is the reduction. Three different types of lactonases acting on sugar derivatives have been characterized in mammalian tissues: 6-phosphogluconolactonase, uronolactonase and aldonolactonase. The first one is an enzyme of the pentose phosphate pathway, which belongs, in mammals, to the same family of proteins as glucosamine 6-phosphate isomerase [44] and has no (direct) role to play in the formation of vitamin C. Uronolactonase, a microsomal enzyme, hydrolyzes d-glucurono-3,6-lactone (K_m≈ 8 mm), but is inactive against aldonolactones [45]. It is a metal-dependent enzyme, but its sequence is presently unknown.

Aldonolactonase (gulonolactonase) is also a metal-dependent enzyme, acting best with Mn²⁺, which is present in the cytosol and hydrolyzes a number of γ- and δ-lactones of a variety of 5-, 6-, and 7-carbon aldonates, including l-gulono-1,4-lactone and d-glucono-1,5-lactone [45]. It also catalyzes the lactonization of aldonates, e.g. of l-gulonate, and can therefore participate in the formation of vitamin C [46]. It has recently been identified as SMP30 (senescence marker protein 30) [47], also known as regucalcin, a protein homologous to bacterial gluconolactonases [48], and which was initially thought to regulate liver cell functions related to Ca²⁺[49] and to be possibly involved in senescence because of its decreased expression with age in liver, kidney and lung [50,51]. The finding that targeted inactivation of the SMP30 gene leads to vitamin C deficiency [47] strongly argues in favour of the involvement of gulonolactonase in the ascorbate synthesis pathway in mammals. This conclusion is in line with earlier findings indicating that l-gulonate rather than d-glucuronolactone is an intermediate in this pathway, such as the fact that ascorbate is more readily formed from d-glucuronate than from d-glucuronolactone in liver extracts [52], and with the lower K_m of glucuronate reductase for d-glucuronate than for d-glucuronolactone (see above).

L-Gulonolactone oxidase

Characterization of the enzyme and of the catalyzed reaction

GLO, a microsomal enzyme, catalyzes aerobically the conversion of l-gulonolactone to l-ascorbate with production of H₂O₂[53,54]. The immediate oxidation product of GLO is 2-keto-l-gulonolactone, an intermediate that spontaneously isomerizes to l-ascorbate [54].

The preferred substrate of the enzyme is l-gulono-1,4-lactone, but it also acts on l-galactono-, d-mannono- and d-altrono-1,4-lactone [55]. Other γ-lactones, including l-idono- and d-gluconolactone, were not oxidized by the enzyme, indicating its configurational specificity for the hydroxyl group at C2. K_m values for l-gulonolactone ranging from 0.007 to 0.15 mm have been reported [55,56]. The enzyme transfers electrons not only to O₂, but also to artificial electron acceptors such as phenazine methosulfate and ferricyanide, although not to cytochrome b₅ and cytochrome c[57]. The production of H₂O₂ is unusual for an enzyme of the endoplasmic reticulum, and one may wonder if this membrane-bound oxidoreductase does not transfer its electrons to another acceptor in intact cells, most particularly because its plant homologues do so. The latter share ∼30% sequence identity with mammalian GLO and differ from this enzyme in three main aspects: (1) they act specifically on l-galactono-1,4-lactone [58,59], which is their physiological substrate; (2) they are bound to the inner mitochondrial membrane [60]; and (3) they do not transfer electrons directly to O₂, but to cytochrome c[59].

GLO is a 50.6 kDa protein [61], which, as indicated by sequence comparisons, is related to plant l-galactonolactone dehydrogenase and fungal d-arabinonolactone oxidase, and more distantly to 6-hydroxynicotine oxidase, d-2-hydroxyglutarate dehydrogenase and 24-dehydrocholesterol reductase, all flavin adenine dinucleotide (FAD)-linked enzymes. Each monomer of GLO binds one molecule of FAD, which is covalently linked to a histidyl residue [55,62]. This residue is presumably His54, which aligns with the histidine (His72) that covalently links FAD in Arthrobacter nicotinovorans 6-hydroxy-d-nicotine oxidase, as indicated by inspection of the structure (pdb file 2BVH) of this bacterial enzyme.

The requirement of detergents for the solubilization of GLO from the microsomal fraction [55–57] strongly suggests its membrane localization. Accordingly, the amino acid sequence of the protein contains several strongly hydrophobic regions, which are thus possibly associated with the endoplasmic reticulum membrane [61]. These regions are predicted to form β-sheets rather than the typical transmembrane helical structure. The orientation of the catalytic site towards the lumen of the endoplasmic reticulum is indicated by the intraluminal accumulation of ascorbate and the preferential intraluminal glutathione oxidation (presumably by hydrogen peroxide) in rat liver microsomes incubated with gulonolactone [63]. It should be noted that the GLO sequence is apparently devoid of targeting motifs for the endoplasmic reticulum, as indicated by analysis of the sequence with the TargetP program [64].

Molecular defects in man and other species

Early enzymological studies identified GLO deficiency as the reason for the inability of some species such as man and guinea pig to synthesize their own vitamin C [65]. Man [66] and guinea pig [67] both have a gene homologous to the rat GLO gene, but they are highly mutated. Compared with the rat gene, which comprises 12 exons, two coding exons (I and V) are missing in its guinea pig homologue [67].

Nucleotide sequence alignment of one exon of the GLO gene from rat with the corresponding exon in the highly mutated, nonfunctional GLO gene of several primates revealed that nucleotide substitutions have occurred at random throughout the primate sequence, as expected for the exon of a gene that ceased to be active during evolution and subsequently evolved without functional constraint [68]. From these two examples, and the finding that ascorbate-deficient species are also observed in several other lineages, it appears that inactivation of the GLO gene occurred several times during evolution, suggesting that the loss of this gene may be advantageous to some species. It has been proposed that the formation of hydrogen peroxide by GLO and the glutathione depletion that ensues are detrimental [69] and that the selective pressure to keep the ability of forming vitamin C is lost in species with ample dietary supply of ascorbic acid.

Rat, pig and mouse models of vitamin C deficiency are known. The osteogenic disorder Shionogi rat is a mutant rat of the Wistar strain deficient in GLO activity and thus unable to synthesize ascorbate. The GLO cDNA of this mutant was found to contain a single base mutation leading to a Cys→Tyr substitution at position 61 in the amino acid sequence [70]. Overexpression experiments in COS-1 cells suggested that this mutation leads to instability of the mutant GLO protein and is responsible for the enzymatic defect in the osteogenic disorder Shionogi rat. The latter manifests deformity, shortening of the legs, multiple fractures, osteoporosis, growth retardation and haemorrhagic tendency when it is fed an ascorbate-deficient diet; these symptoms are largely prevented by providing vitamin C in the food [71]. A similar symptomatology is found in the od/od pig, in which the GLO gene is inactivated due to an intragenic deletion removing exon 8 [72].

GLO knockout mice have also been generated through homologous recombination and the effects of vitamin C deficiency in these mice have been studied [73]. The mutant mice depend on dietary vitamin C supplementation for survival. The most striking effects of a low vitamin C diet on the knockout mice were alterations in their aortic walls (for instance fragmentation of elastic lamina), which were proposed to be caused by defects in the crosslinking of collagen and elastin.

An alternative fate for glucuronate: the pentose pathway

As described above, d-glucuronate can be metabolized to ascorbate in most vertebrates, well known exceptions being primates and the guinea pig. By contrast, in all animal species examined, glucuronate can be converted to the pentose l-xylulose in a pathway known as the pentose pathway or the glucuronic acid oxidation pathway [74]. The reactions involved in this pathway are represented in Fig. 2. The reduction step catalyzed by glucuronate reductase (see previous section) is shared by the ascorbate synthesis pathway and the pentose pathway.

In the latter, l-gulonate is then oxidized to 3-keto-l-gulonate by an NAD-dependent dehydrogenase [75]. The cDNA encoding rabbit liver l-gulonate 3-dehydrogenase has recently been cloned [76]. The enzyme, shown to be identical to lens λ-crystallin, displays a K_m of ∼0.2 mm for l-gulonate. 3-Keto-l-gulonate is decarboxylated to l-xylulose by a poorly characterized decarboxylase [77] whose molecular identity is unknown. l-Xylulose is then converted to xylitol by an NADPH-dependent l-xylulose reductase. A cDNA encoding a dicarbonyl/l-xylulose reductase has been cloned from a mouse kidney cDNA library [78]. This reductase displays a marked preference for NADPH over NADH and is ubiquitously expressed in several mammalian species. A K_m value of 0.21 mm for l-xylulose has been reported for the human recombinant enzyme.

Xylitol is oxidized to d-xylulose by an NAD-dependent enzyme identical to sorbitol dehydrogenase [79]. Finally, d-xylulose can enter the pentose phosphate pathway after its phosphorylation by d-xylulokinase. The latter has been purified to homogeneity from bovine liver and was shown to be a monomeric enzyme of 51 kDa [80]. The pure enzyme acted on d-xylulose and d-ribulose with respective K_m values of 0.14 and 0.27 mm. A human cDNA encoding a ‘xylulokinase-like’ protein of 528 amino acids has been isolated [81]. The predicted gene product bears 22% identity to the xylulokinase of Haemophilus influenzae.

The occurrence of the pentose pathway in humans is indicated by the fact that rare individuals excrete abnormal quantities of l-xylulose (1–4 g·day⁻¹) in the urine. This benign condition, known as essential pentosuria [74], was recognized by Garrod, almost a century ago, as an inborn error of metabolism. In 1929, Margolis [82] noted that ingestion of aminopyrine leads to a marked increase in pentose excretion in pentosuric subjects and some years later it was shown that this effect could be mimicked by the administration, not only of a series of other drugs, but also of glucuronic acid [74]. This effect, which is very reminiscent of the stimulation exerted by nonglucuronidable hydrophobic drugs on vitamin C formation in animals [28], is most probably also due to a stimulation of the UDP-glucuronidase activity of UGTs.

Pentosuria, an autosomal recessive trait, is due to l-xylulose reductase deficiency [83]. Lane [84] reported the separation of a major and a minor isozyme for l-xylulose reductase in human erythrocytes. In pentosuric subjects, only the minor isozyme, which displayed a K_m for l-xylulose of ∼100 mm, was detected upon electrophoresis and ion-exchange chromatography. This suggests that homozygosity for the pentosuria allele results in deficiency of the major isozyme, which most probably corresponds to the recently cloned dicarbonyl/l-xylulose reductase (see above). The genetic defect underlying pentosuria has not yet been reported.

The benign nature of this condition (the only symptom is the elevated urinary pentose excretion) shows that the pentose pathway does not play an indispensable role in human metabolism. While in most mammalian species, this pathway produces a precursor for the formation of ascorbate (l-gulonate), in humans and some other species it probably essentially allows the return of a portion of glucuronate carbon to mainstream carbohydrate metabolism.

Control of vitamin C synthesis

Outline on the regulation of vitamin C synthesis

The main control is apparently exerted at the level of the formation of glucuronate from UDP-glucuronate, as enhancement of this conversion is accompanied by an increase in the formation of vitamin C [28]. However, the pathway is branched at the level of l-gulonate and the proportion of l-gulonate that is converted to vitamin C or l-xylulose must depend on the relative activities of the rate-limiting enzymes downstream in the pathways. In the case of vitamin C formation, the rate-limiting step downstream is catalyzed by l-gulonolactone oxidase, as indicated by the observation that heterozygous (OD/od) pigs for GLO deficiency have (when fed an ascorbate-free diet) a plasma ascorbic acid level amounting to ∼50% of that found in control (OD/OD) pigs [72]. A comparison between the amount of d-glucuronate that accumulates in isolated rat hepatocytes incubated with various xenobiotics in the presence of the glucuronate reductase inhibitor sorbinil (vitamin C formation is then blocked!) and the amount of ascorbate that is formed under similar conditions but in the absence of sorbinil indicates that ∼30% of l-gulonate is directed towards ascorbate formation [28].

Effect of xenobiotics

As mentioned above, the stimulatory effect of xenobiotics has been ascribed, at least partially, to a short-term effect on UDP-glucuronidase, i.e. most likely UGTs. How the various (always nonglucuronidable) xenobiotics act is still unknown. Their important structural diversity and their hydrophobic character suggest that they could act by perturbing locally the phospholipidic environment of UGTs and by thus inducing a conformational change favouring a hydrolase activity of these transferases. Alternatively, these compounds, which are hydrophobic but lack a suitable glucuronosyl acceptor function, could stimulate the hydrolase activity through a pseudosubstrate mechanism. Besides this short-term action, the effect of some xenobiotics to stimulate the expression of UGTs [18,24,25] or GLO [85] is also conducive to a stimulated formation of ascorbic acid.

One may wonder what advantage organisms may derive from the stimulation of vitamin C biosynthesis caused by nonglucuronidable xenobiotics. There is no answer at present to this question, but an interesting possibility would be that the stimulatory xenobiotics are membrane-perturbing agents which could favour the generation of reactive oxygen species when inserted in membranes with active electron transport, the increased vitamin C availability playing then a protective role.

Effect of glutathione

From a quantitative point of view, glutathione and vitamin C are the most abundant reducing agents in cells. Furthermore, GSH is implicated in vitamin C recycling from DHA (see next section). It would therefore make sense for glutathione to exert a control on vitamin C synthesis. Several experiments performed with glutathione-depleting agents indicate that glutathione depletion favours vitamin C synthesis. Administration to adult mice of buthionine sulfoximine, an inhibitor of glutathione synthesis, led to a two-fold increase in the amount of vitamin C in liver within 4 h [86]. Similarly, incubation of rat hepatocytes with 1-bromoheptane or phorone, which are conjugated with GSH, caused a more than two-fold increase in vitamin C content after 2 h of incubation [87]. On the basis of the observation that a series of glutathione-depleting agents including, surprisingly, dibutyryl cyclic AMP, enhanced vitamin C formation and also glycogenolysis in murine hepatocytes, it was proposed that increased ascorbate synthesis is the result of a ‘push effect’ involving an increase in the concentration of UDP-glucose [88]. However, no measurements of UDP-glucose or UDP-glucuronate were made to substantiate this hypothesis. Furthermore, the potent glycogenolytic agent glucagon does not stimulate glucuronate [28] or vitamin C [87] formation in rat hepatocytes, and the effects of some of the compounds that were tested by Braun et al. [88] could not be reproduced by other authors [28,87].

The mechanism of the effect of glutathione-depleting agents is therefore presently not understood. One may wonder to what extent some of the agents used to deplete glutathione do not act like xenobiotics, by stimulating the formation of glucuronate from UDP-glucuronate through a direct action on UDP-glucuronidase. A more exciting possibility would be that the control is exerted downstream on GLO. If this were the case, glutathione depletion should enhance the conversion of l-gulonolactone to vitamin C. Finally, one has also to consider the theoretical possibility that the glutathione-depleting agents might act by slowing down vitamin C degradation.

Recycling of vitamin C

As described in the introduction, ascorbate plays major roles as a water-soluble antioxidant and as a cofactor of several enzymes, which lead to its one-electron oxidation to SDA. Disproportionation of SDA (2 SDA→ascorbate + DHA), in turn, results in the formation of DHA. Both SDA and DHA are reduced back to ascorbate by several enzymatic systems that are briefly reviewed below and schematized in Fig. 3.

Reduction of semidehydroascorbate

The reduction of SDA in the cytosol has been assigned to enzymes using NADH (NADH-cytochrome b₅ reductase) or NADPH (thioredoxin reductase). SDA can also be reduced in the lumen of neuroendocrine secretory vesicles or in the external medium by transmembrane electron transfer systems.

NADH-cytochrome b₅ reductase

Early studies showed that oxidation of NADH by liver microsomes was stimulated in the presence of SDA, and microsomal NADH-cytochrome b₅ reductase was proposed to participate in the electron transfer system, since the purified enzyme itself was able to reduce SDA in the presence of NADH [89]. Ito et al. [90] showed that, in rat liver, most of the NADH-dependent SDA reductase activity is localized in the outer mitochondrial membrane. Some activity could be detected in the nuclear and microsomal fractions. Inhibition of the mitochondrial SDA reductase activity with specific antibodies suggested participation of NADH-cytochrome b₅ reductase and of a cytochrome b₅-like protein of the outer mitochondrial membrane [90].

NADH-cytochrome b₅ reductase is an FAD-containing enzyme [91,92], which exists as a 300 amino acid membrane-bound form and a 275 amino acid soluble form [93]. The membrane-bound protein is located mainly in the endoplasmic reticulum and the outer mitochondrial membranes, but a small fraction of the enzyme is apparently also associated with the plasma membrane [94]. Its C-terminal catalytic domain (∼275 amino acid residues) is oriented towards the cytosol [95]. The soluble form (identical to the catalytic domain of the membrane-bound form) of the protein is found mainly in erythrocytes, where it is involved in the reduction of methaemoglobin [95]. Fibroblasts of a patient with methaemoglobinaemia due to a mutation in the NADH-cytochrome b₅ reductase gene were shown to be deficient in NADH-dependent SDA reductase activity [96], which confirms the involvement of this enzyme in SDA reduction.

An NADH-linked soluble SDA reductase was also purified from rabbit lens, but the N-terminal sequence of a peptide fragment prepared from this protein did not show significant similarity with any known protein sequence [97]. This suggests that, besides NADH-cytochrome b₅ reductase, additional NADH-dependent enzymes may participate in SDA reduction.

Thioredoxin reductase

Purified rat liver thioredoxin reductase, a selenoprotein, was shown to catalyze the NADPH-dependent reduction of SDA with a K_m value (in the presence of thioredoxin, which acts as an activator) of ∼3 µm for this radical [98]. NADPH-dependent reduction of SDA was also demonstrated in dialyzed cytosolic fractions prepared from rat liver, where it was found to be enhanced by selenocystine and inhibited by aurothioglucose (an inhibitor of selenoenzymes), indicating that it is contributed by thioredoxin reductase [98]. This interpretation was further supported by the finding that the NADPH-dependent SDA reduction was markedly decreased (by ∼75%) in liver cytosolic fractions derived from selenium-deficient rats, which have low (<10% of control rats) thioredoxin reductase activity.

Cytochrome b₅₆₁-mediated reduction of intravesicular SDA

Ascorbate is a cofactor of dopamine β-hydroxylase and of peptidylglycine α-amidating monooxygenase (see Introduction). Both enzymes are localized in neuroendocrine secretory vesicles (catecholamine-storing vesicles for the first enzyme and peptide-storing vesicles for the second) and catalyze reactions generating SDA [99]. As ascorbate and SDA do not cross the membrane of the storage vesicles, the reduction of SDA to ascorbate occurs inside the vesicles and involves the transmembrane transfer of electrons by cytochrome b₅₆₁, the second most abundant protein in the chromaffin granule membrane [100,101]. Electrons are furnished to cytochrome b₅₆₁ by cytosolic ascorbate, which is maintained under its reduced form by the mitochondrial outer membrane SDA reductase [102]. The intravesicular reduction of SDA to ascorbate is thermodynamically driven by the pH gradient (SDA reduction involves the binding of a proton and is therefore favoured at acidic pH) and the membrane potential (positive inside) created across the chromaffin vesicle membrane by an inwardly directed proton-translocating ATPase [103]. The sequence of cytochrome b₅₆₁ is known [104], but this protein has not yet been crystallized. Current structural models suggest that cytochrome b₅₆₁ forms six transmembrane α-helices and contains two hemes with differing redox potentials, each anchored by a pair of well-conserved histidyl residues, one near the cytosolic and the other near the intravesicular face of the membrane [105–107].

Reduction of extracellular SDA involving transfer of electrons across the plasma membrane

Erythrocytes are able to transfer electrons from intracellular donors to extracellular SDA. In a system where SDA was generated extracellularly from ascorbate by ascorbate oxidase, it was shown that erythrocytes decreased both the ascorbate oxidation rate and the steady-state extracellular SDA concentration [108]. Using a similar experimental model, Van Duijn et al. [109] showed that erythrocytes prevented the degradation of extracellular ascorbate by reactions that were driven by intracellular ascorbate and NADH. The relative contributions of endogenous ascorbate and NADH could not be established, but the results clearly indicated that intracellular ascorbate donates electrons to extracellular SDA via a plasma membrane redox system. The electron transport system still needs to be identified. It does not involve cytochrome b₅₆₁, which is absent from the erythrocyte plasma membrane [110].

Isolated rat liver plasma membranes catalyze SDA reduction in the presence of NADH and this activity is inhibited by lectins [111], indicating the involvement of glycoprotein(s). Furthermore, HL-60 cells slow down the (chemically induced) oxidation of external ascorbate [112]. This effect is increased when NADH formation is stimulated by the addition of lactate to the cells and is inhibited by lectins. These and other observations suggest the existence of an NADH-dependent trans-plasma membrane redox system reducing SDA to ascorbate in several cell types [113], although part of the observed stabilization of external ascorbate by cells could be explained by transition ion chelation and thus inhibition of ascorbate autoxidation [114,115]. The enzymatic system seems to be distinct from the plasma membrane NADH-dependent ferricyanide reductase [116] and to involve coenzyme Q [117], and NADH-cytochrome b₅ reductase [118], as well as other, as yet unidentified, factors [119]. A system allowing the utilization of reducing equivalents in the extracellular medium (experimentally, by ferricyanide) derived from intracellular ascorbate appears also to be present in HL-60 cells [120].

Reduction of dehydroascorbate

This process is probably less important quantitatively than the reduction of SDA, which is the most important oxidation product of ascorbate. Intracellular DHA may, however, derive from SDA through disproportionation or through import from extracellular sources.

Spontaneous reaction with GSH

DHA can be nonenzymatically reduced to ascorbate by GSH [121]. This reaction, which presumably proceeds through a glutathione-ascorbate thiohemiketal adduct, is thermodynamically favourable (ΔE°′ =0.16 V; K_eq ≈ 2.5 × 10⁵ m⁻¹ at pH 7), but is relatively slow at physiological concentrations of GSH. From the data reported by Winkler et al. [121], it can be calculated that at 5 mm GSH and 0.1 mm DHA (pH 7.0; 20–22 °C), this reaction would be half-complete after ∼10 min, which compares with a half-life of ∼100 min for the spontaneous hydrolysis of DHA to 2,3-diketogulonate at the same temperature [122]. However, as described in the next section, the hydrolysis of DHA is enhanced by bicarbonate and by an intracellular lactonase. These considerations lead to the conclusion that an important (irreversible) loss of vitamin C would occur if the spontaneous reaction with GSH were solely responsible for the reduction of DHA to ascorbate. However, as detailed below, several enzymes facilitate this reaction with GSH. Furthermore, NADPH-dependent reductases appear also to be involved in DHA reduction.

Glutaredoxin and protein disulfide isomerase

Highly purified preparations of glutaredoxin (also known as thioltransferase) from pig liver, beef thymus and human placenta, as well as of bovine liver protein disulfide isomerase (PDI) display DHA reductase activity [123]. Glutaredoxin and PDI are thiol-disulfide oxidoreductases, which both possess active-site cysteine residues. Glutaredoxins are small (12 kDa) cytosolic enzymes that physiologically use GSH as a reductant, whereas PDI is a larger (57 kDa) protein, bound to the endoplasmic reticulum membrane, which serves to create and rearrange disulfide bonds [123]. Kinetic analysis of their DHA reductase activity revealed K_m values in the range of 0.2–2.2 mm and 1.6–8.7 mm for DHA and GSH, respectively. Based on the subcellular localization of these two types of enzymes, it was proposed that glutaredoxin could contribute to ascorbate recycling from DHA in the cytosol, whereas PDI might catalyze this reaction in the endoplasmic reticulum. The finding that PDI can act on DHA led to the proposal that the latter could be implicated as an oxidant in protein disulfide formation catalyzed by PDI [124,125].

More recently, experiments with cultured human lens epithelial cells in which thioltransferase (glutaredoxin) was overexpressed or knocked down, indicated that this enzyme plays a major role in ascorbate recycling in these cells [126]. Other studies have shown that PDI is intrinsically much poorer (∼250-fold lower catalytic efficiency) as a DHA reductase than glutaredoxin [127].

Omega class glutathione transferase

Maellaro et al. [128] purified a cytosolic rat liver DHA reductase, which was clearly different from glutaredoxin and PDI. SDS/PAGE analysis gave a single band at 31 kDa and K_m values of 0.25 and 2.8 mm were reported for DHA and GSH, respectively. The enzyme had a specific requirement for GSH as hydrogen donor. Immunotitration experiments with polyclonal antibodies revealed that it accounted for ∼70% of the GSH-dependent DHA reductase activity of a rat liver cytosol [129]. Other studies with the same enzyme purified from human erythrocytes showed it to have a catalytic efficiency comparable to that of glutaredoxin in its DHA reductase activity (k_cat/K_m ratio of ∼ 25 000 m⁻¹·s⁻¹ in both cases) [127]. The enzyme was found to be broadly distributed among organs, as revealed by activity measurement, immunoblot analysis [129] and northern blot analysis [130].

The cDNA of rat liver GSH-dependent DHA reductase has been cloned and found to encode a protein with a predicted molecular weight of ∼25 kDa [130]. DHA reductase is not homologous to glutaredoxin or PDI, but belongs to the superfamily of glutathione S-transferases (GSTs), more precisely to the Omega class of GSTs (GSTO; this class of GSTs has recently been reviewed in [131]). GSTs generally catalyze the transfer of GSH to a variety of substrates including aromatic compounds and leukotrienes. The crystal structure of human GSTO1, the orthologue of rat GSH-dependent DHA reductase, shows the presence in the active site of a cysteine (Cys32), conserved in GSTOs, at a position where GSTs from other classes contain a tyrosine or a serine residue serving to stabilize bound GSH as a thiolate via the hydroxyl group [131,132]. In contrast, Cys32 of GSTO1 was found to form a disulfide bond with GSH [132].

GSTOs have very low glutathione conjugating activity, but they act as thioltransferases, and as reductases for DHA and monomethylarsonate (an intermediate in the arsenic biotransformation pathway) [131]. Concerning DHA reduction, human GSTO2 (which shares ∼64% identity with human GSTO1, but has a more restricted tissue distribution) was found to be 70–100 times more active than GSTO1 [133]. The role of GSH-dependent DHA reductase in the recycling of vitamin C was substantiated by showing that CHO cells expressing rat liver DHA reductase accumulated more ascorbate than nontransfected cells when incubated in the presence of DHA [130].

3α-Hydroxysteroid dehydrogenase

Del Bello et al. [134] purified an NADPH-dependent DHA reductase from rat liver cytosol. This enzyme had, however, a lower affinity for DHA (K_m = 4.6 mm) than the GSH-dependent enzymes mentioned above. The monomeric structure, the relatively low molecular weight (∼37 kDa), the NADPH-dependence, as well as the cytosolic localization of the enzyme suggested that it belongs to the aldo-keto reductase superfamily. The substrate specificity as well as the inhibition pattern of the enzyme (reduction of 5α-androstane-3,17-dione and inhibition by steroidal and nonsteroidal anti-inflammatory drugs) pointed to a possible identity with 3α-hydroxysteroid dehydrogenase, which was finally proven by microsequence analysis.

Thioredoxin reductase

In addition to its activity on SDA (see previous subsection), purified rat liver thioredoxin reductase was also shown to catalyze the NADPH-dependent reduction of DHA [135]. The K_m value for this substrate (0.7 mm), though >200-fold higher than that for SDA (3 µm), is in the same range as the K_m values of the GSH-dependent DHA reductases. Arguments mentioned above to support the role of thioredoxin reductase in the reduction of SDA (decreased NADPH-dependent reductase activity in liver cytosolic fractions of selenium-deficient rats; inhibition by aurothioglucose) also apply to the reduction of DHA and indicate that thioredoxin reductase is responsible for at least 75% of the NADPH-dependent reduction of DHA in a rat liver cytosol. It should be mentioned that GSH-dependent DHA reductase activity in cytosolic fractions was found to be typically two- to three-fold higher than the total NADPH-dependent reductase activity and that it was not affected by selenium deficiency [135].

Assessment of the role of the reductive systems

As mentioned above, SDA can be either directly reduced in a one-electron step to ascorbate or further oxidized (e.g. by disproportionation) to DHA before the latter is reduced in a two-electron step. Coassin et al. [136] compared the NADH-dependent SDA reductase and the GSH-dependent DHA reductase activities in homogenates of pig tissues and concluded that, at the low physiological concentrations (micromolar range) of DHA, recycling of ascorbate is effectively accomplished by reduction of the ascorbyl radical, DHA reduction occurring only at a negligible rate under these conditions. No DHA reductase activity was detected at 0.1 mm DHA. Reduction of DHA is, however, certainly important when this compound is imported from the extracellular medium.

Studies in which glutathione was depleted have pointed to the importance of the GSH-dependent reduction of DHA. Administration of l-buthionine-(S,R)-sulfoximine (BSO), a transition-state inactivator of γ-glutamylcysteine synthetase, to newborn rats decreased tissue levels of GSH, but also of ascorbate and markedly increased DHA [137]. Severe tissue damage was observed especially at the level of mitochondria and the animals died within a few days. Simultaneous administration of ascorbate decreased their mortality and increased glutathione levels. Newborn rats given lower doses of BSO developed cataracts. This could also be partially prevented by the administration of ascorbate. These in vivo observations showed that one of the roles of glutathione is to maintain ascorbate in its reduced state and that ascorbate can partially replace glutathione in some of its antioxidant functions. Adult rats and mice are much less sensitive to BSO treatment than newborn animals. Mortality was not increased by BSO administration to adult mice and much less severe tissue damage was observed [86]. The higher sensitivity of newborn animals was proposed to be due to lower ascorbate synthesis in the first days of life, but may also partly be explained by the higher retention of BSO in tissues of newborn animals than of adult animals [137]. These and other studies emphasizing an in vivo relationship between glutathione and ascorbate in animals that can or cannot synthesize ascorbate have been reviewed by Meister [138].

Catabolism of vitamin C

In mammals, the degradation of ascorbate appears to proceed via DHA, an unstable molecule, and to involve spontaneous and possibly enzyme-catalyzed reactions. A specific pathway for the degradation of vitamin C has only been fully described in bacteria.

Spontaneous breakdown of dehydroascorbate

Ascorbate is oxidized to DHA in a variety of enzymatic and nonenzymatic reactions (see Introduction). DHA can be reduced back to ascorbate (see previous section), but can also be hydrolyzed to 2,3-diketo-l-gulonate (2,3-DKG). The latter reaction is irreversible and 2,3-DKG, unlike DHA, is therefore devoid of antiscorbutic activity. DHA is unstable and nonenzymatic hydrolysis occurs rather rapidly at neutral pH, the half-time of decay being about 5–15 min at 37 °C [122]. Incubation of DHA or 2,3-DKG in the presence of phosphate buffer (pH 7) for several hours at 37 °C leads to the formation of l-erythrulose and oxalate [139], indicating cleavage between the second and the third carbon (Fig. 4). When the incubation is performed in the presence of H₂O₂ (which may be formed during chemical oxidation of ascorbate), l-threonate is the major degradation product formed (Fig. 4) and no l-erythrulose is detected [139]. A five-carbon intermediate (3,4,5-trihydroxy-2-ketopentanoate) is also transiently formed in the presence of H₂O₂[140], indicating that the reaction proceeds by successive cleavage of the bonds between C1 and C2, and C2 and C3.

It was speculated that the highly reactive ketose, l-erythrulose, if formed in vivo, could play a role in ascorbate-dependent modifications of protein observed in vitro and proposed to occur in vivo in human lens during diabetic and age-onset cataract formation [139].

Degradation of vitamin C in mammals

In vivo studies

In human subjects injected with ascorbate labelled on C1, an average of 44% of the total radioactivity excreted in urine was recovered as oxalate; other urinary metabolites were 2,3-DKG (∼20%) and DHA (< 2%), and ∼20% of the total urinary radioactivity was present as ascorbate [141]. Little if any radioactivity was recovered as CO₂. These findings indicate that a major catabolic event in man is the cleavage of the C6 molecule (presumably a spontaneous cleavage of 2,3-DKG) between C2 and C3, with little if any decarboxylation. The oxalate formed in this way may contribute to the formation of kidney stones in susceptible individuals. However, the association between ascorbate supplementation and increased risk of kidney stone formation remains a matter of controversy [142].

The global human metabolism of ascorbic acid differs from that of animal species in several respects [141,143]. Firstly, the half-life of ascorbate was shown to be ∼4–5 times longer in man than in guinea pigs and rats. Second, in guinea pigs and rats, radiolabelled CO₂ is produced from ascorbate labelled on C1. Third, whereas in humans DHA is almost completely converted to ascorbate, this process is less efficient in other animals where DHA is rapidly catabolized. Finally, on vitamin C-free diet, the onset of scurvy is less rapid in man than in guinea pigs. These findings indicate that man is better able than rodents to save vitamin C, probably because of both a better ability to reduce DHA to ascorbate and a lower dehydroascorbatase activity (see below). We speculate that this also applies to bats, which are unable to synthesize ascorbic acid and some of which have a low supply of this vitamin in their food [13].

In vitro studies

As mentioned above, DHA is unstable and its nonenzymatic hydrolysis occurs rather rapidly at pH 7.0 [122]. It is accelerated by bicarbonate, which is responsible for the rapid disappearance of DHA in blood plasma [144]. An enzyme catalyzing the delactonization of DHA has been detected and partially purified from beef liver [143,145]. This enzyme hydrolyzes a series of other lactones and shares many properties with gulonolactonase, from which it could not be separated [143]. Lower activities of both dehydroascorbatase [145] and gulonolactonase (aldonolactonase) [45,146] are present in tissues of primates compared to other mammals. Assuming that both activities are contributed by the same enzyme, this lower activity may be beneficial in primates because it would slow down the irreversible loss of DHA while not having any detrimental effect on ascorbate formation (which is absent). It would be interesting to know if the bone loss observed in transgenic rats overexpressing regucalcin [147,148] (which, as noted above, has recently been identified as the aldonolactonase involved in ascorbate synthesis [47]) is related to vitamin C deficiency.

As for the formation of oxalate, which is one of the major excretion products of ascorbate in humans, no enzyme has yet been described that catalyzes the cleavage of 2,3-DKG between the second and the third carbon. It is therefore likely that this reaction takes place spontaneously. The metabolic fate of the other product (most probably l-erythrulose, since H₂O₂ levels are kept low) is unknown. The finding that addition of ascorbate or DHA to glycogen-depleted mouse hepatocytes causes an increase in the formation of glucose and in the concentration of d-xylulose 5-phosphate [149] suggests that they can be converted to a metabolizable sugar or sugar derivative.

Utilization of vitamin C by Escherichia coli

E. coli can ferment ascorbate and the operon responsible for ascorbate utilization has recently been identified [150]. This identification was based on the hypothesis that the catabolic pathway of ascorbate could lead to the formation of the pentose phosphate intermediate d-xylulose 5-phosphate, via 4-epimerization of l-ribulose 5-phosphate. The latter reaction is also involved in the catabolic pathway of l-arabinose, where it is catalyzed by AraD. Two multigene operons of unknown function were found to comprise an AraD homologue in the genome of E. coli K-12.

The deletion of one of these operons generated mutants that failed to ferment ascorbate [150]. The ‘utilization of l-ascorbate’ operon (ula operon) thus identified contains six genes (designated ulaA, B, C, D, E and F) (Fig. 5A). UlaA, UlaB and UlaC are components of a PTS system involved in the uptake of ascorbate and its phosphorylation to l-ascorbate 6-phosphate [151]. The proteins encoded by ulaD, ulaE and ulaF were functionally characterized as 3-keto-l-gulonate 6-phosphate decarboxylase, l-xylulose 5-phosphate 3-epimerase and l-ribulose 5-phosphate 4-epimerase, respectively [150]. A gene located upstream of the ula operon, ulaG, encodes a distant homologue of a metal-dependent hydrolase that is essential for ascorbate utilization [151]. It was proposed to encode a cytoplasmic ascorbate 6-phosphate lactonase catalyzing the conversion of ascorbate 6-phosphate to 3-keto-l-gulonate 6-phosphate.

All together the divergently transcribed operons ulaABCDEF and ulaG encode proteins allowing the uptake of ascorbate and its conversion to d-xylulose 5-phosphate and CO₂ (Fig. 5C). They are both regulated by the nearby ulaR gene and together constitute a regulon [152]. ulaR encodes a repressor belonging to the DeoR family of bacterial regulatory proteins and inactivation of this gene leads to constitutive expression of the ula operons. A detailed study of the regulation of expression of the ula operons by UlaR and other factors has been reported [153].

Disruption of the other operon (yia) containing an AraD homologue (Fig. 5B) did not affect the fermentation of ascorbate [150]. This operon encodes nine different proteins, five of which have been overexpressed and characterized [150]. On this basis, it has been proposed that the yia operon allows the metabolism of 2,3-diketo-l-gulonate through a pathway in which it is successively reduced to 3-keto-l-gulonate by YiaK (a dehydrogenase belonging to a novel family of pyridine nucleotide-linked oxidoreductases [154]) and phosphorylated to 3-keto-l-gulonate 6-phosphate by an ATP-dependent kinase (YiaP) (Fig. 5C). 3-Keto-l-gulonate 6-phosphate would then be converted to d-xylulose 5-phosphate in a sequence of reactions similar to that described above for the ula operon. The gene products involved are YiaQ (3-keto-l-gulonate 6-phosphate decarboxylase), YiaR (l-xylulose 5-phosphate 3-epimerase) and YiaS (l-ribulose 5-phosphate 4-epimerase). It should be noted that no enzymatic activity could be detected in the case of YiaR despite its high degree of identity with UlaE (l-xylulose 5-phosphate 3-epimerase) [150]. The operon also encodes three proteins of a putative ABC transport system (YiaM, YiaN and YiaO), possibly involved in the transport of 2,3-DKG [150]. However, the definitive proof that this operon is involved in the utilization of the hydrolyzed form of DHA is still lacking.

A search for similar operons in other bacterial genomes using the SEED database (http://theseed.uchicago.edu) indicates that the first (ula) occurs not only in Enterobacteriae (E. coli, Shigella dysenteriae, Shigella sonnei, Shigella flexneri, Salmonella enterica, Salmonella paratyphi, Salmonella typhimurium, and Klebsiella pneumoniae), but also in Lactobacillales (Enterococcus faecium, Streptococcus agalactiae, Streptococcus pneumoniae, Streptococcus pyogenes, and Streptococcus uberis) and in Mycoplasma penetrans, Mycoplasma pneumoniae, and Mycoplasma synoviae. The second type of operon (yia) appears to be much less frequent. We noted its presence only in E. coli, S. enterica, S. typhimurium, and Serratia marcescens. Mammalian genomes apparently do not encode an orthologue of 3-keto-l-gulonate 6-phosphate decarboxylase, the critical enzyme in these vitamin C degradation pathways, as indicated by blast searches.

Acknowledgements

This work was supported by the Concerted Research Action Program of the Communauté Française de Belgique; the Interuniversity Attraction Poles Program, Belgian Science Policy; and the Fonds de la Recherche Scientifique Médicale. CLL was a fellow of the Fonds National de la Recherche Scientifique (FNRS).

References

1 Padayatty SJ, Katz A, Wang Y, Eck P, Kwon O, Lee JH, Chen S, Corpe C, Dutta A, Dutta SK & Levine M (2003) Vitamin C as an antioxidant: evaluation of its role in disease prevention. J Am Coll Nutr 22, 18–35.
10.1080/07315724.2003.10719272
CASPubMedWeb of Science®Google Scholar
2 Buettner GR (1993) The pecking order of free radicals and antioxidants: lipid peroxidation, alpha-tocopherol, and ascorbate. Arch Biochem Biophys 300, 535–543.
10.1006/abbi.1993.1074
CASPubMedWeb of Science®Google Scholar
3 Buettner GR & Jurkiewicz BA (1996) Catalytic metals, ascorbate and free radicals: combinations to avoid. Radiat Res 145, 532–541.
10.2307/3579271
CASPubMedWeb of Science®Google Scholar
4 Prigge ST, Mains RE, Eipper BA & Amzel LM (2000) New insights into copper monooxygenases and peptide amidation: structure, mechanism and function. Cell Mol Life Sci 57, 1236–1259.
10.1007/PL00000763
CASPubMedWeb of Science®Google Scholar
5 Englard S & Seifter S (1986) The biochemical functions of ascorbic acid. Annu Rev Nutr 6, 365–406.
10.1146/annurev.nu.06.070186.002053
CASPubMedWeb of Science®Google Scholar
6 Hirota K & Semenza GL (2005) Regulation of hypoxia-inducible factor 1 by prolyl and asparaginyl hydroxylases. Biochem Biophys Res Commun 338, 610–616.
10.1016/j.bbrc.2005.08.193
CASPubMedWeb of Science®Google Scholar
7 Wilson JX (2005) Regulation of vitamin C transport. Annu Rev Nutr 25, 105–125.
10.1146/annurev.nutr.25.050304.092647
CASPubMedWeb of Science®Google Scholar
8 Takanaga H, Mackenzie B & Hediger MA (2004) Sodium-dependent ascorbic acid transporter family SLC23. Pflugers Arch 447, 677–682.
10.1007/s00424-003-1104-1
CASPubMedWeb of Science®Google Scholar
9 Liang WJ, Johnson D & Jarvis SM (2001) Vitamin C transport systems of mammalian cells. Mol Membr Biol 18, 87–95.
10.1080/09687680110033774
CASPubMedWeb of Science®Google Scholar
10 Sotiriou S, Gispert S, Cheng J, Wang Y, Chen A, Hoogstraten-Miller S, Miller GF, Kwon O, Levine M, Guttentag SH et al. (2002) Ascorbic-acid transporter Slc23a1 is essential for vitamin C transport into the brain and for perinatal survival. Nat Med 8, 514–517.
10.1038/0502-514
CASPubMedWeb of Science®Google Scholar
11 Moreau R & Dabrowski K (1998) Body pool and synthesis of ascorbic acid in adult sea lamprey (Petromyzon marinus): an agnathan fish with gulonolactone oxidase activity. Proc Natl Acad Sci USA 95, 10279–10282.
10.1073/pnas.95.17.10279
CASPubMedWeb of Science®Google Scholar
12 Moreau R & Dabrowski K (1998) Fish acquired ascorbic acid synthesis prior to terrestrial vertebrate emergence. Free Radic Biol Med 25, 989–990.

CASPubMedWeb of Science®Google Scholar
13 Birney EC, Jenness R & Ayaz KM (1976) Inability of bats to synthesise l-ascorbic acid. Nature 260, 626–628.
10.1038/260626a0
CASPubMedWeb of Science®Google Scholar
14 Chatterjee IB (1973) Evolution and the biosynthesis of ascorbic acid. Science 182, 1271–1272.
10.1126/science.182.4118.1271
CASPubMedWeb of Science®Google Scholar
15 Smirnoff N, Conklin PL & Loewus FA (2001) Biosynthesis of ascorbic acid in plants: a renaissance. Annu Rev Plant Physiol Plant Mol Biol 52, 437–467.
10.1146/annurev.arplant.52.1.437
CASPubMedWeb of Science®Google Scholar
16 Smirnoff N (2001) l-Ascorbic acid biosynthesis. Vitam Horm 61, 241–266.
10.1016/S0083-6729(01)61008-2
CASPubMedWeb of Science®Google Scholar
17 Longenecker HE, Fricke HH & King CG (1940) The effect of organic compounds upon vitamin C synthesis in the rat. J Biol Chem 135, 497–510.
10.1016/S0021-9258(18)73119-0
CASWeb of Science®Google Scholar
18 Hollmann S & Touster O (1962) Alterations in tissue levels of uridine diphosphate glucose dehydrogenase, uridine diphosphate glucuronic acid pyrophosphatase and glucuronyl transferase induced by substances influencing the production of ascorbic acid. Biochim Biophys Acta 62, 338–352.
10.1016/0006-3002(62)90048-3
CASPubMedWeb of Science®Google Scholar
19 Horio F & Yoshida A (1982) Effects of some xenobiotics on ascorbic acid metabolism in rats. J Nutr 112, 416–425.
10.1093/jn/112.3.416
CASPubMedWeb of Science®Google Scholar
20 Burns JJ, Mosbach EH & Schulenberg S (1954) Ascorbic acid synthesis in normal and drug-treated rats, studied with l-ascorbic-1-C¹⁴ acid. J Biol Chem 207, 679–687.
10.1016/S0021-9258(18)65685-6
CASPubMedWeb of Science®Google Scholar
21 Burns JJ, Evans C & Trousof N (1957) Stimulatory effect of barbital on urinary excretion of l-ascorbic acid and nonconjugated d-glucuronic acid. J Biol Chem 227, 785–794.
10.1016/S0021-9258(18)70758-8
CASPubMedWeb of Science®Google Scholar
22 Evans C, Conney AH, Trousof N & Burns JJ (1960) Metabolism of d-galactose to d-glucuronic acid, l-gulonic acid and l-ascorbic acid in normal and barbital-treated rats. Biochim Biophys Acta 41, 9–14.
10.1016/0006-3002(60)90362-0
CASPubMedWeb of Science®Google Scholar
23 Burns JJ & Evans C (1956) The synthesis of l-ascorbic acid in the rat from d-glucuronolactone and l-gulonolactone. J Biol Chem 223, 897–905.
10.1016/S0021-9258(18)65088-4
CASPubMedWeb of Science®Google Scholar
24 Horio F, Kimura M & Yoshida A (1983) Effect of several xenobiotics on the activities of enzymes affecting ascorbic acid synthesis in rats. J Nutr Sci Vitaminol (Tokyo) 29, 233–247.
10.3177/jnsv.29.233
CASPubMedWeb of Science®Google Scholar
25 Horio F, Shibata T, Makino S, Machino S, Hayashi Y, Hattori T & Yoshida A (1993) UDP glucuronosyltransferase gene expression is involved in the stimulation of ascorbic acid biosynthesis by xenobiotics in rats. J Nutr 123, 2075–2084.

CASPubMedWeb of Science®Google Scholar
26 Iyanagi T, Watanabe T & Uchiyama Y (1989) The 3-methylcholanthrene-inducible UDP-glucuronosyltransferase deficiency in the hyperbilirubinemic rat (Gunn rat) is caused by a -1 frameshift mutation. J Biol Chem 264, 21302–21307.
10.1016/S0021-9258(19)30079-1
CASPubMedWeb of Science®Google Scholar
27 Iyanagi T (1991) Molecular basis of multiple UDP-glucuronosyltransferase isoenzyme deficiencies in the hyperbilirubinemic rat (Gunn rat). J Biol Chem 266, 24048–24052.
10.1016/S0021-9258(18)54390-8
CASPubMedWeb of Science®Google Scholar
28 Linster CL & Van Schaftingen E (2003) Rapid stimulation of free glucuronate formation by non-glucuronidable xenobiotics in isolated rat hepatocytes. J Biol Chem 278, 36328–36333.
10.1074/jbc.M306593200
CASPubMedWeb of Science®Google Scholar
29 Ginsburg V, Weissbach A & Maxwell ES (1958) Formation of glucuronic acid from uridinediphosphate glucuronic acid. Biochim Biophys Acta 28, 649–650.
10.1016/0006-3002(58)90538-9
CASPubMedWeb of Science®Google Scholar
30 Puhakainen E & Hänninen O (1976) Pyrophosphatase and glucuronosyltransferase in microsomal UDPglucuronic-acid metabolism in the rat liver. Eur J Biochem 61, 165–169.
10.1111/j.1432-1033.1976.tb10007.x
CASPubMedWeb of Science®Google Scholar
31 Pogell BM & Leloir LF (1961) Nucleotide activation of liver microsomal glucuronidation. J Biol Chem 236, 293–298.
10.1016/S0021-9258(18)64355-8
CASPubMedWeb of Science®Google Scholar
32 Linster CL & Van Schaftingen E (2006) Glucuronate, the precursor of vitamin C, is directly formed from UDP-glucuronate in liver. FEBS J 273, 1516–1527.
10.1111/j.1742-4658.2006.05172.x
CASPubMedWeb of Science®Google Scholar
33 Bossuyt X & Blanckaert N (1995) Mechanism of stimulation of microsomal UDP-glucuronosyltransferase by UDP-N-acetylglucosamine. Biochem J 305, 321–328.
10.1042/bj3050321
PubMedWeb of Science®Google Scholar
34 Evans WH (1974) Nucleotide pyrophosphatase, a sialoglycoprotein located on the hepatocyte surface. Nature 250, 391–394.
10.1038/250391a0
CASPubMedWeb of Science®Google Scholar
35 Touster O, Aronson NN Jr, Dulaney JT & Hendrickson H (1970) Isolation of rat liver plasma membranes. Use of nucleotide pyrophosphatase and phosphodiesterase I as marker enzymes. J Cell Biol 47, 604–618.
10.1083/jcb.47.3.604
CASPubMedWeb of Science®Google Scholar
36 Hochman Y & Zakim D (1984) Studies of the catalytic mechanism of microsomal UDP-glucuronyltransferase. Alpha-glucuronidase activity and its stimulation by phospholipids. J Biol Chem 259, 5521–5525.
10.1016/S0021-9258(18)91043-4
CASPubMedWeb of Science®Google Scholar
37 Mano Y, Suzuki K, Yamada K & Shimazono N (1961) Enzymic studies on TPN l-hexonate dehydrogenase from rat liver. J Biochem (Tokyo) 49, 618–634.
10.1093/oxfordjournals.jbchem.a127352
CASPubMedWeb of Science®Google Scholar
38 Jez JM & Penning TM (2001) The aldo-keto reductase (AKR) superfamily: an update. Chem Biol Interact 130–132, 499–525.
10.1016/S0009-2797(00)00295-7
CASPubMedWeb of Science®Google Scholar
39 Bohren KM, Bullock B, Wermuth B & Gabbay KH (1989) The aldo-keto reductase superfamily. cDNAs and deduced amino acid sequences of human aldehyde and aldose reductases. J Biol Chem 264, 9547–9551.
10.1016/S0021-9258(18)60566-6
CASPubMedWeb of Science®Google Scholar
40 Griffin BW (1992) Functional and structural relationships among aldose reductase, l-hexonate dehydrogenase (aldehyde reductase), and recently identified homologous proteins. Enzyme Microb Technol 14, 690–695.
10.1016/0141-0229(92)90107-Y
CASPubMedWeb of Science®Google Scholar
41 Srivastava SK, Ansari NH, Hair GA & Das B (1984) Aldose and aldehyde reductases in human tissues. Biochim Biophys Acta 800, 220–227.
10.1016/0304-4165(84)90399-4
CASPubMedWeb of Science®Google Scholar
42 Petrash JM & Srivastava SK (1982) Purification and properties of human liver aldehyde reductases. Biochim Biophys Acta 707, 105–114.
10.1016/0167-4838(82)90402-2
CASPubMedWeb of Science®Google Scholar
43 Bhatnagar A, Liu S, Das B, Ansari NH & Srivastava SK (1990) Inhibition kinetics of human kidney aldose and aldehyde reductases by aldose reductase inhibitors. Biochem Pharmacol 39, 1115–1124.
10.1016/0006-2952(90)90292-S
CASPubMedWeb of Science®Google Scholar
44 Collard F, Collet JF, Gerin I, Veiga-da-Cunha M & Van Schaftingen E (1999) Identification of the cDNA encoding human 6-phosphogluconolactonase, the enzyme catalyzing the second step of the pentose phosphate pathway. FEBS Lett 459, 223–226.
10.1016/S0014-5793(99)01247-8
CASPubMedWeb of Science®Google Scholar
45 Winkelman J & Lehninger AL (1958) Aldono- and uronolactonases of animal tissues. J Biol Chem 233, 794–799.
10.1016/S0021-9258(18)64657-5
CASPubMedWeb of Science®Google Scholar
46 Bublitz C & Lehninger AL (1961) The role of aldonolactonase in the conversion of l-gulonate to l-ascorbate. Biochim Biophys Acta 47, 288–297.
10.1016/0006-3002(61)90289-X
CASWeb of Science®Google Scholar
47 Kondo Y, Inai Y, Sato Y, Handa S, Kubo S, Shimokado K, Goto S, Nishikimi M, Maruyama N & Ishigami A (2006) Senescence marker protein 30 functions as gluconolactonase in l-ascorbic acid biosynthesis, and its knockout mice are prone to scurvy. Proc Natl Acad Sci USA 103, 5723–5728.
10.1073/pnas.0511225103
CASPubMedWeb of Science®Google Scholar
48 Kanagasundaram V & Scopes R (1992) Isolation and characterization of the gene encoding gluconolactonase from Zymomonas mobilis. Biochim Biophys Acta 1171, 198–200.
10.1016/0167-4781(92)90120-O
CASPubMedWeb of Science®Google Scholar
49 Yamaguchi M (2005) Role of regucalcin in maintaining cell homeostasis and function (review). Int J Mol Med 15, 371–389.

CASPubMedWeb of Science®Google Scholar
50 Fujita T, Uchida K & Maruyama N (1992) Purification of senescence marker protein-30 (SMP30) and its androgen-independent decrease with age in the rat liver. Biochim Biophys Acta 1116, 122–128.
10.1016/0304-4165(92)90108-7
CASPubMedWeb of Science®Google Scholar
51 Mori T, Ishigami A, Seyama K, Onai R, Kubo S, Shimizu K, Maruyama N & Fukuchi Y (2004) Senescence marker protein-30 knockout mouse as a novel murine model of senile lung. Pathol Int 54, 167–173.
10.1111/j.1440-1827.2003.01603.x
CASPubMedWeb of Science®Google Scholar
52 Stirpe F & Comporti M (1965) Regulation of ascorbic acid and of xylulose synthesis in rat-liver extracts. The effect of starvation on the enzymes of the glucuronic acid pathway. Biochem J 95, 354–362.
10.1042/bj0950354
CASPubMedWeb of Science®Google Scholar
53 Chatterjee IB, Chatterjee GC, Ghosh NC, Ghosh JJ & Guha BC (1960) Biological synthesis of l-ascorbic acid in animal tissues: conversion of l-gulonolactone into l-ascorbic acid. Biochem J 74, 193–203.
10.1042/bj0740193
CASPubMedWeb of Science®Google Scholar
54 Chatterjee IB, Chatterjee GC, Ghosh NC, Ghosh JJ & Guha BC (1960) Biological synthesis of l-ascorbic acid in animal tissues: conversion of d-glucuronolactone and l-gulonolactone into l-ascorbic acid. Biochem J 76, 279–292.
10.1042/bj0760279
CASPubMedWeb of Science®Google Scholar
55 Kiuchi K, Nishikimi M & Yagi K (1982) Purification and characterization of l-gulonolactone oxidase from chicken kidney microsomes. Biochemistry 21, 5076–5082.
10.1021/bi00263a035
CASPubMedWeb of Science®Google Scholar
56 Nishikimi M, Tolbert BM & Udenfriend S (1976) Purification and characterization of l-gulono-gamma-lactone oxidase from rat and goat liver. Arch Biochem Biophys 175, 427–435.
10.1016/0003-9861(76)90530-0
CASPubMedWeb of Science®Google Scholar
57 Eliceiri GL, Lai EK & McCay PB (1969) Gulonolactone oxidase. Solubilization, properties, and partial purification. J Biol Chem 244, 2641–2645.
10.1016/S0021-9258(18)83447-0
CASPubMedWeb of Science®Google Scholar
58 Ôba K, Ishikawa S, Nishikawa M, Mizuno H & Yamamoto T (1995) Purification and properties of l-galactono-gamma-lactone dehydrogenase, a key enzyme for ascorbic acid biosynthesis, from sweet potato roots. J Biochem (Tokyo) 117, 120–124.
10.1093/oxfordjournals.jbchem.a124697
CASPubMedWeb of Science®Google Scholar
59 Østergaard J, Persiau G, Davey MW, Bauw G & Van Montagu M (1997) Isolation of a cDNA coding for l-galactono-gamma-lactone dehydrogenase, an enzyme involved in the biosynthesis of ascorbic acid in plants. Purification, characterization, cDNA cloning, and expression in yeast. J Biol Chem 272, 30009–30016.
10.1074/jbc.272.48.30009
CASPubMedWeb of Science®Google Scholar
60 Siendones E, González-Reyes JA, Santos-Ocaña C, Navas P & Córdoba F (1999) Biosynthesis of ascorbic acid in kidney bean. l-galactono-gamma-lactone dehydrogenase is an intrinsic protein located at the mitochondrial inner membrane. Plant Physiol 120, 907–912.
10.1104/pp.120.3.907
CASPubMedWeb of Science®Google Scholar
61 Koshizaka T, Nishikimi M, Ozawa T & Yagi K (1988) Isolation and sequence analysis of a complementary DNA encoding rat liver l-gulono-gamma-lactone oxidase, a key enzyme for l-ascorbic acid biosynthesis. J Biol Chem 263, 1619–1621.
10.1016/S0021-9258(19)77923-X
CASPubMedWeb of Science®Google Scholar
62 Kenney WC, Edmondson DE & Singer TP (1976) Identification of the covalently bound flavin of l-gulono-gamma-lactone oxidase. Biochem Biophys Res Commun 71, 1194–1200.
10.1016/0006-291X(76)90780-4
CASPubMedWeb of Science®Google Scholar
63 Puskás F, Braun L, Csala M, Kardon T, Marcolongo P, Benedetti A, Mandl J & Bánhegyi G (1998) Gulonolactone oxidase activity-dependent intravesicular glutathione oxidation in rat liver microsomes. FEBS Lett 430, 293–296.
10.1016/S0014-5793(98)00678-4
CASPubMedWeb of Science®Google Scholar
64 Emanuelsson O, Nielsen H, Brunak S & von Heijne G (2000) Predicting subcellular localization of proteins based on their N-terminal amino acid sequence. J Mol Biol 300, 1005–1016.
10.1006/jmbi.2000.3903
CASPubMedWeb of Science®Google Scholar
65 Burns JJ, Peyser P & Moltz A (1956) Missing step in guinea pigs required for the biosynthesis of l-ascorbic acid. Science 124, 1148–1149.
10.1126/science.124.3232.1148-a
CASPubMedWeb of Science®Google Scholar
66 Nishikimi M, Fukuyama R, Minoshima S, Shimizu N & Yagi K (1994) Cloning and chromosomal mapping of the human nonfunctional gene for l-gulono-gamma-lactone oxidase, the enzyme for l-ascorbic acid biosynthesis missing in man. J Biol Chem 269, 13685–13688.
10.1016/S0021-9258(17)36884-9
CASPubMedWeb of Science®Google Scholar
67 Nishikimi M, Kawai T & Yagi K (1992) Guinea pigs possess a highly mutated gene for l-gulono-gamma-lactone oxidase, the key enzyme for l-ascorbic acid biosynthesis missing in this species. J Biol Chem 267, 21967–21972.
10.1016/S0021-9258(19)36707-9
CASPubMedWeb of Science®Google Scholar
68 Ohta Y & Nishikimi M (1999) Random nucleotide substitutions in primate nonfunctional gene for l-gulono-gamma-lactone oxidase, the missing enzyme in l-ascorbic acid biosynthesis. Biochim Biophys Acta 1472, 408–411.
10.1016/S0304-4165(99)00123-3
CASPubMedWeb of Science®Google Scholar
69 Bánhegyi G, Csala M, Braun L, Garzó T & Mandl J (1996) Ascorbate synthesis-dependent glutathione consumption in mouse liver. FEBS Lett 381, 39–41.
10.1016/0014-5793(96)00077-4
CASPubMedWeb of Science®Google Scholar
70 Kawai T, Nishikimi M, Ozawa T & Yagi K (1992) A missense mutation of l-gulono-gamma-lactone oxidase causes the inability of scurvy-prone osteogenic disorder rats to synthesize l-ascorbic acid. J Biol Chem 267, 21973–21976.
10.1016/S0021-9258(19)36708-0
CASPubMedWeb of Science®Google Scholar
71 Mizushima Y, Harauchi T, Yoshizaki T & Makino S (1984) A rat mutant unable to synthesize vitamin C. Experientia 40, 359–361.
10.1007/BF01952551
CASPubMedWeb of Science®Google Scholar
72 Hasan L, Vögeli P, Stoll P, Kramer ŠŠ, Stranzinger G & Neuenschwander S (2004) Intragenic deletion in the gene encoding l-gulonolactone oxidase causes vitamin C deficiency in pigs. Mamm Genome 15, 323–333.
10.1007/s00335-003-2324-6
CASPubMedGoogle Scholar
73 Maeda N, Hagihara H, Nakata Y, Hiller S, Wilder J & Reddick R (2000) Aortic wall damage in mice unable to synthesize ascorbic acid. Proc Natl Acad Sci USA 97, 841–846.
10.1073/pnas.97.2.841
CASPubMedWeb of Science®Google Scholar
74 Hiatt HH (2001) Pentosuria. In The Metabolic and Molecular Bases of Inherited Disease ( CR Scriver, AL Beaudet, WS Sly & D Valle, eds), 8th edn, Vol. I, pp. 1589–1599. McGraw-Hill, New York.

Google Scholar
75 Smiley JD & Ashwell G (1961) Purification and properties of beta-l-hydroxy acid dehydrogenase. II. Isolation of beta-keto-l-gulonic acid, an intermediate in l-xylulose biosynthesis. J Biol Chem 236, 357–364.
10.1016/S0021-9258(18)64367-4
CASWeb of Science®Google Scholar
76 Ishikura S, Usami N, Araki M & Hara A (2005) Structural and functional characterization of rabbit and human l-gulonate 3-dehydrogenase. J Biochem (Tokyo) 137, 303–314.
10.1093/jb/mvi033
CASPubMedGoogle Scholar
77 Winkelman J & Ashwell G (1961) Enzymic formation of l-xylulose from beta-keto-l-gulonic acid. Biochim Biophys Acta 52, 170–175.
10.1016/0006-3002(61)90914-3
CASPubMedGoogle Scholar
78 Nakagawa J, Ishikura S, Asami J, Isaji T, Usami N, Hara A, Sakurai T, Tsuritani K, Oda K, Takahashi M et al. (2002) Molecular characterization of mammalian dicarbonyl/l-xylulose reductase and its localization in kidney. J Biol Chem 277, 17883–17891.
10.1074/jbc.M110703200
CASPubMedWeb of Science®Google Scholar
79 Marini I, Bucchioni L, Borella P, Del Corso A & Mura U (1997) Sorbitol dehydrogenase from bovine lens: purification and properties. Arch Biochem Biophys 340, 383–391.
10.1006/abbi.1997.9882
CASPubMedWeb of Science®Google Scholar
80 Dills WL Jr, Parsons PD, Westgate CL & Komplin NJ (1994) Assay, purification, and properties of bovine liver d-xylulokinase. Protein Expr Purif 5, 259–265.
10.1006/prep.1994.1039
CASPubMedWeb of Science®Google Scholar
81 Tamari M, Daigo Y, Ishikawa S & Nakamura Y (1998) Genomic structure of a novel human gene (XYLB) on chromosome 3p22→p21.3 encoding a xylulokinase-like protein. Cytogenet Cell Genet 82, 101–104.
10.1159/000015076
CASPubMedWeb of Science®Google Scholar
82 Margolis JI (1929) Chronic pentosuria and migraine. Am J Med Sci 177, 348–371.
10.1097/00000441-192903000-00005
Google Scholar
83 Wang YM & Van Eys J (1970) The enzymatic defect in essential pentosuria. N Engl J Med 282, 892–896.
10.1056/NEJM197004162821604
CASPubMedWeb of Science®Google Scholar
84 Lane AB (1985) On the nature of l-xylulose reductase deficiency in essential pentosuria. Biochem Genet 23, 61–72.
10.1007/BF00499113
CASPubMedWeb of Science®Google Scholar
85 Braun L, Mile V, Schaff Z, Csala M, Kardon T, Mandl J & Bánhegyi G (1999) Induction and peroxisomal appearance of gulonolactone oxidase upon clofibrate treatment in mouse liver. FEBS Lett 458, 359–362.
10.1016/S0014-5793(99)01184-9
CASPubMedWeb of Science®Google Scholar
86 Mårtensson J & Meister A (1992) Glutathione deficiency increases hepatic ascorbic acid synthesis in adult mice. Proc Natl Acad Sci USA 89, 11566–11568.
10.1073/pnas.89.23.11566
CASPubMedWeb of Science®Google Scholar
87 Chan TS, Wilson JX & O'Brien PJ (2004) Glycogenolysis is directed towards ascorbate synthesis by glutathione conjugation. Biochem Biophys Res Commun 317, 149–156.
10.1016/j.bbrc.2004.03.022
CASPubMedWeb of Science®Google Scholar
88 Braun L, Csala M, Poussu A, Garzó T, Mandl J & Bánhegyi G (1996) Glutathione depletion induces glycogenolysis dependent ascorbate synthesis in isolated murine hepatocytes. FEBS Lett 388, 173–176.
10.1016/0014-5793(96)00548-0
CASPubMedWeb of Science®Google Scholar
89 Iyanagi T & Yamazaki I (1969) One-electron-transfer reactions in biochemical systems. III. One-electron reduction of quinones by microsomal flavin enzymes. Biochim Biophys Acta 172, 370–381.
10.1016/0005-2728(69)90133-9
CASPubMedWeb of Science®Google Scholar
90 Ito A, Hayashi S & Yoshida T (1981) Participation of a cytochrome b₅-like hemoprotein of outer mitochondrial membrane (OM cytochrome b) in NADH-semidehydroascorbic acid reductase activity of rat liver. Biochem Biophys Res Commun 101, 591–598.
10.1016/0006-291X(81)91300-0
CASPubMedWeb of Science®Google Scholar
91 Takano T, Ogawa K, Sato M, Bando S & Yubisui T (1987) Preliminary X-ray data of NADH-cytochrome b₅ reductase from human erythrocytes. J Mol Biol 195, 749–750.
10.1016/0022-2836(87)90195-1
CASPubMedWeb of Science®Google Scholar
92 Shirabe K, Yubisui T & Takeshita M (1989) Expression of human erythrocyte NADH-cytochrome b₅ reductase as an alpha-thrombin-cleavable fused protein in Escherichia coli. Biochim Biophys Acta 1008, 189–192.
10.1016/0167-4781(80)90007-X
CASPubMedWeb of Science®Google Scholar
93 Percy MJ, McFerran NV & Lappin TR (2005) Disorders of oxidised haemoglobin. Blood Rev 19, 61–68.
10.1016/j.blre.2004.02.001
CASPubMedWeb of Science®Google Scholar
94 Borgese N & Pietrini G (1986) Distribution of the integral membrane protein NADH-cytochrome b₅ reductase in rat liver cells, studied with a quantitative radioimmunoblotting assay. Biochem J 239, 393–403.
10.1042/bj2390393
CASPubMedWeb of Science®Google Scholar
95 Borgese N, D'Arrigo A, De Silvestris M & Pietrini G (1993) NADH-cytochrome b₅ reductase and cytochrome b₅ isoforms as models for the study of post-translational targeting to the endoplasmic reticulum. FEBS Lett 325, 70–75.
10.1016/0014-5793(93)81416-W
CASPubMedWeb of Science®Google Scholar
96 Shirabe K, Landi MT, Takeshita M, Uziel G, Fedrizzi E & Borgese N (1995) A novel point mutation in a 3′ splice site of the NADH-cytochrome b₅ reductase gene results in immunologically undetectable enzyme and impaired NADH-dependent ascorbate regeneration in cultured fibroblasts of a patient with type II hereditary methemoglobinemia. Am J Hum Genet 57, 302–310.

CASPubMedWeb of Science®Google Scholar
97 Bando M, Inoue T, Oka M, Nakamura K, Kawai K, Obazawa H, Kobayashi S & Takehana M (2004) Isolation of ascorbate free radical reductase from rabbit lens soluble fraction. Exp Eye Res 79, 869–873.
10.1016/j.exer.2004.08.011
CASPubMedGoogle Scholar
98 May JM, Cobb CE, Mendiratta S, Hill KE & Burk RF (1998) Reduction of the ascorbyl free radical to ascorbate by thioredoxin reductase. J Biol Chem 273, 23039–23045.
10.1074/jbc.273.36.23039
CASPubMedWeb of Science®Google Scholar
99 Njus D & Kelley PM (1993) The secretory-vesicle ascorbate-regenerating system: a chain of concerted H⁺/e^–-transfer reactions. Biochim Biophys Acta 1144, 235–248.
10.1016/0005-2728(93)90108-R
CASPubMedWeb of Science®Google Scholar
100 Njus D, Knoth J, Cook C & Kelley PM (1983) Electron transfer across the chromaffin granule membrane. J Biol Chem 258, 27–30.
10.1016/S0021-9258(18)33212-5
CASPubMedWeb of Science®Google Scholar
101 Fleming PJ & Kent UM (1991) Cytochrome b₅₆₁, ascorbic acid, and transmembrane electron transfer. Am J Clin Nutr 54, 1173S–1178S.
10.1093/ajcn/54.6.1173s
CASPubMedWeb of Science®Google Scholar
102 Diliberto EJ Jr, Dean G, Carter C & Allen PL (1982) Tissue, subcellular, and submitochondrial distributions of semidehydroascorbate reductase: possible role of semidehydroascorbate reductase in cofactor regeneration. J Neurochem 39, 563–568.
10.1111/j.1471-4159.1982.tb03982.x
CASPubMedWeb of Science®Google Scholar
103 Njus D, Kelley PM, Harnadek GJ & Pacquing YV (1987) Mechanism of ascorbic acid regeneration mediated by cytochrome b₅₆₁. Ann NY Acad Sci 493, 108–119.
10.1111/j.1749-6632.1987.tb27188.x
CASPubMedWeb of Science®Google Scholar
104 Perin MS, Fried VA, Slaughter CA & Südhof TC (1988) The structure of cytochrome b₅₆₁, a secretory vesicle-specific electron transport protein. EMBO J 7, 2697–2703.
10.1002/j.1460-2075.1988.tb03123.x
CASPubMedWeb of Science®Google Scholar
105 Bashtovyy D, Bérczi A, Asard H & Páli T (2003) Structure prediction for the di-heme cytochrome b₅₆₁ protein family. Protoplasma 221, 31–40.
10.1007/s00709-002-0065-0
CASPubMedWeb of Science®Google Scholar
106 Bérczi A, Su D, Lakshminarasimhan M, Vargas A & Asard H (2005) Heterologous expression and site-directed mutagenesis of an ascorbate-reducible cytochrome b₅₆₁. Arch Biochem Biophys 443, 82–92.
10.1016/j.abb.2005.09.006
CASPubMedWeb of Science®Google Scholar
107 Takigami T, Takeuchi F, Nakagawa M, Hase T & Tsubaki M (2003) Stopped-flow analyses on the reaction of ascorbate with cytochrome b₅₆₁ purified from bovine chromaffin vesicle membranes. Biochemistry 42, 8110–8118.
10.1021/bi0267588
CASPubMedWeb of Science®Google Scholar
108 May JM, Qu Z & Cobb CE (2000) Extracellular reduction of the ascorbate free radical by human erythrocytes. Biochem Biophys Res Commun 267, 118–123.
10.1006/bbrc.1999.1906
CASPubMedWeb of Science®Google Scholar
109 Van Duijn MM, Tijssen K, VanSteveninck J, Van den Broek PJ & Van der Zee J (2000) Erythrocytes reduce extracellular ascorbate free radicals using intracellular ascorbate as an electron donor. J Biol Chem 275, 27720–27725.
10.1074/jbc.M910281199
CASPubMedWeb of Science®Google Scholar
110 Van Duijn MM, Buijs JT, Van der Zee J & Van den Broek PJ (2001) The ascorbate: ascorbate free radical oxidoreductase from the erythrocyte membrane is not cytochrome b₅₆₁. Protoplasma 217, 94–100.
10.1007/BF01289418
CASPubMedGoogle Scholar
111 Navas P, Estévez A, Burón MI, Villalba JM & Crane FL (1988) Cell surface glycoconjugates control the activity of the NADH-ascorbate free radical reductase of rat liver plasma membrane. Biochem Biophys Res Commun 154, 1029–1033.
10.1016/0006-291X(88)90243-4
CASPubMedWeb of Science®Google Scholar
112 Alcain FJ, Buron MI, Villalba JM & Navas P (1991) Ascorbate is regenerated by HL-60 cells through the transplasmalemma redox system. Biochim Biophys Acta 1073, 380–385.
10.1016/0304-4165(91)90146-8
CASPubMedWeb of Science®Google Scholar
113 May JM (1999) Is ascorbic acid an antioxidant for the plasma membrane? FASEB J 13, 995–1006.
10.1096/fasebj.13.9.995
CASPubMedWeb of Science®Google Scholar
114 Schweinzer E, Waeg G, Esterbauer H & Goldenberg H (1993) No enzymatic activities are necessary for the stabilization of ascorbic acid by K-562 cells. FEBS Lett 334, 106–108.
10.1016/0014-5793(93)81691-R
CASPubMedWeb of Science®Google Scholar
115 Rodríguez-Aguilera JC & Navas P (1994) Extracellular ascorbate stabilization: enzymatic or chemical process? J Bioenerg Biomembr 26, 379–384.
10.1007/BF00762778
CASPubMedWeb of Science®Google Scholar
116 Villalba JM, Canalejo A, Rodríguez-Aguilera JC, Burón MI, Morré DJ & Navas P (1993) NADH-ascorbate free radical and -ferricyanide reductase activities represent different levels of plasma membrane electron transport. J Bioenerg Biomembr 25, 411–417.
10.1007/BF00762467
CASPubMedWeb of Science®Google Scholar
117 Villalba JM, Navarro F, Córdoba F, Serrano A, Arroyo A, Crane FL & Navas P (1995) Coenzyme Q reductase from liver plasma membrane: purification and role in trans-plasma-membrane electron transport. Proc Natl Acad Sci USA 92, 4887–4891.
10.1073/pnas.92.11.4887
CASPubMedWeb of Science®Google Scholar
118 Navarro F, Villalba JM, Crane FL, Mackellar WC & Navas P (1995) A phospholipid-dependent NADH-coenzyme Q reductase from liver plasma membrane. Biochem Biophys Res Commun 212, 138–143.
10.1006/bbrc.1995.1947
CASPubMedWeb of Science®Google Scholar
119 Arroyo A, Rodríguez-Aguilera JC, Santos-Ocaña C, Villalba JM & Navas P (2004) Stabilization of extracellular ascorbate mediated by coenzyme Q transmembrane electron transport. Methods Enzymol 378, 207–217.
10.1016/S0076-6879(04)78017-9
CASPubMedGoogle Scholar
120 Van Duijn MM, Van der Zee J, VanSteveninck J & Van den Broek PJ (1998) Ascorbate stimulates ferricyanide reduction in HL-60 cells through a mechanism distinct from the NADH-dependent plasma membrane reductase. J Biol Chem 273, 13415–13420.
10.1074/jbc.273.22.13415
CASPubMedWeb of Science®Google Scholar
121 Winkler BS, Orselli SM & Rex TS (1994) The redox couple between glutathione and ascorbic acid: a chemical and physiological perspective. Free Radic Biol Med 17, 333–349.
10.1016/0891-5849(94)90019-1
CASPubMedWeb of Science®Google Scholar
122 Bode AM, Cunningham L & Rose RC (1990) Spontaneous decay of oxidized ascorbic acid (dehydro-l-ascorbic acid) evaluated by high-pressure liquid chromatography. Clin Chem 36, 1807–1809.
10.1093/clinchem/36.10.1807
CASPubMedWeb of Science®Google Scholar
123 Wells WW, Xu DP, Yang Y & Rocque PA (1990) Mammalian thioltransferase (glutaredoxin) and protein disulfide isomerase have dehydroascorbate reductase activity. J Biol Chem 265, 15361–15364.
10.1016/S0021-9258(18)55401-6
CASPubMedWeb of Science®Google Scholar
124 Wells WW & Xu DP (1994) Dehydroascorbate reduction. J Bioenerg Biomembr 26, 369–377.
10.1007/BF00762777
CASPubMedWeb of Science®Google Scholar
125 Bánhegyi G, Csala M, Szarka A, Varsányi M, Benedetti A & Mandl J (2003) Role of ascorbate in oxidative protein folding. Biofactors 17, 37–46.
10.1002/biof.5520170105
CASPubMedWeb of Science®Google Scholar
126 Fernando MR, Satake M, Monnier VM & Lou MF (2004) Thioltranferase mediated ascorbate recycling in human lens epithelial cells. Invest Ophthalmol Vis Sci 45, 230–237.
10.1167/iovs.03-0545
PubMedWeb of Science®Google Scholar
127 Xu DP, Washburn MP, Sun GP & Wells WW (1996) Purification and characterization of a glutathione dependent dehydroascorbate reductase from human erythrocytes. Biochem Biophys Res Commun 221, 117–121.
10.1006/bbrc.1996.0555
CASPubMedWeb of Science®Google Scholar
128 Maellaro E, Del Bello B, Sugherini L, Santucci A, Comporti M & Casini AF (1994) Purification and characterization of glutathione-dependent dehydroascorbate reductase from rat liver. Biochem J 301, 471–476.
10.1042/bj3010471
CASPubMedWeb of Science®Google Scholar
129 Paolicchi A, Pezzini A, Saviozzi M, Piaggi S, Andreuccetti M, Chieli E, Malvaldi G & Casini AF (1996) Localization of a GSH-dependent dehydroascorbate reductase in rat tissues and subcellular fractions. Arch Biochem Biophys 333, 489–495.
10.1006/abbi.1996.0419
CASPubMedWeb of Science®Google Scholar
130 Ishikawa T, Casini AF & Nishikimi M (1998) Molecular cloning and functional expression of rat liver glutathione-dependent dehydroascorbate reductase. J Biol Chem 273, 28708–28712.
10.1074/jbc.273.44.28708
CASPubMedWeb of Science®Google Scholar
131 Whitbread AK, Masoumi A, Tetlow N, Schmuck E, Coggan M & Board PG (2005) Characterization of the omega class of glutathione transferases. Methods Enzymol 401, 78–99.
10.1016/S0076-6879(05)01005-0
CASPubMedWeb of Science®Google Scholar
132 Board PG, Coggan M, Chelvanayagam G, Easteal S, Jermiin LS, Schulte GK, Danley DE, Hoth LR, Griffor MC, Kamath AV et al. (2000) Identification, characterization, and crystal structure of the Omega class glutathione transferases. J Biol Chem 275, 24798–24806.
10.1074/jbc.M001706200
CASPubMedWeb of Science®Google Scholar
133 Schmuck EM, Board PG, Whitbread AK, Tetlow N, Cavanaugh JA, Blackburn AC & Masoumi A (2005) Characterization of the monomethylarsonate reductase and dehydroascorbate reductase activities of Omega class glutathione transferase variants: implications for arsenic metabolism and the age-at-onset of Alzheimer's and Parkinson's diseases. Pharmacogenet Genomics 15, 493–501.
10.1097/01.fpc.0000165725.81559.e3
CASPubMedWeb of Science®Google Scholar
134 Del Bello B, Maellaro E, Sugherini L, Santucci A, Comporti M & Casini AF (1994) Purification of NADPH-dependent dehydroascorbate reductase from rat liver and its identification with 3 alpha-hydroxysteroid dehydrogenase. Biochem J 304, 385–390.
10.1042/bj3040385
CASPubMedWeb of Science®Google Scholar
135 May JM, Mendiratta S, Hill KE & Burk RF (1997) Reduction of dehydroascorbate to ascorbate by the selenoenzyme thioredoxin reductase. J Biol Chem 272, 22607–22610.
10.1074/jbc.272.36.22607
CASPubMedWeb of Science®Google Scholar
136 Coassin M, Tomasi A, Vannini V & Ursini F (1991) Enzymatic recycling of oxidized ascorbate in pig heart: one-electron vs two-electron pathway. Arch Biochem Biophys 290, 458–462.
10.1016/0003-9861(91)90566-2
CASPubMedWeb of Science®Google Scholar
137 Mårtensson J & Meister A (1991) Glutathione deficiency decreases tissue ascorbate levels in newborn rats: ascorbate spares glutathione and protects. Proc Natl Acad Sci USA 88, 4656–4660.
10.1073/pnas.88.11.4656
CASPubMedWeb of Science®Google Scholar
138 Meister A (1994) Glutathione-ascorbic acid antioxidant system in animals. J Biol Chem 269, 9397–9400.
10.1016/S0021-9258(17)36891-6
CASPubMedWeb of Science®Google Scholar
139 Simpson GL & Ortwerth BJ (2000) The non-oxidative degradation of ascorbic acid at physiological conditions. Biochim Biophys Acta 1501, 12–24.
10.1016/S0925-4439(00)00009-0
CASPubMedWeb of Science®Google Scholar
140 Deutsch JC (1998) Oxygen-accepting antioxidants which arise during ascorbate oxidation. Anal Biochem 265, 238–245.
10.1006/abio.1998.2940
CASPubMedWeb of Science®Google Scholar
141 Hellman L & Burns JJ (1958) Metabolism of l-ascorbic acid-1-C¹⁴ in man. J Biol Chem 230, 923–930.
10.1016/S0021-9258(18)70515-2
CASPubMedWeb of Science®Google Scholar
142 Massey LK, Liebman M & Kynast-Gales SA (2005) Ascorbate increases human oxaluria and kidney stone risk. J Nutr 135, 1673–1677.
10.1093/jn/135.7.1673
CASPubMedWeb of Science®Google Scholar
143 Kagawa Y & Takiguchi H (1962) Enzymatic studies on ascorbic acid catabolism in animals. II. Delactonization of dehydro-l-ascorbic acid. J Biochem (Tokyo) 51, 197–203.
10.1093/oxfordjournals.jbchem.a127521
CASPubMedWeb of Science®Google Scholar
144 Koshiishi I, Mamura Y & Imanari T (1998) Bicarbonate promotes a cleavage of lactone ring of dehydroascorbate. Biochim Biophys Acta 1379, 257–263.
10.1016/S0304-4165(97)00106-2
CASPubMedWeb of Science®Google Scholar
145 Kagawa Y, Takiguchi H & Shimazono N (1961) Enzymic delactonization of dehydro-l-ascorbate in animal tissues. Biochim Biophys Acta 51, 413–415.
10.1016/0006-3002(61)90196-2
CASPubMedWeb of Science®Google Scholar
146 Yamada K, Ishikawa S & Shimazono N (1959) On the microsomal and soluble lactonases. Biochim Biophys Acta 32, 253–255.
10.1016/0006-3002(59)90576-1
PubMedWeb of Science®Google Scholar
147 Yamaguchi M, Misawa H, Uchiyama S, Morooka Y & Tsurusaki Y (2002) Role of endogenous regucalcin in bone metabolism: bone loss is induced in regucalcin transgenic rats. Int J Mol Med 10, 377–383.

CASPubMedWeb of Science®Google Scholar
148 Uchiyama S & Yamaguchi M (2004) Bone loss in regucalcin transgenic rats: enhancement of osteoclastic cell formation from bone marrow of rats with increasing age. Int J Mol Med 14, 451–455.

CASPubMedWeb of Science®Google Scholar
149 Braun L, Puskás F, Csala M, Györffy E, Garzó T, Mandl J & Bánhegyi G (1996) Gluconeogenesis from ascorbic acid: ascorbate recycling in isolated murine hepatocytes. FEBS Lett 390, 183–186.
10.1016/0014-5793(96)00654-0
CASPubMedWeb of Science®Google Scholar
150 Yew WS & Gerlt JA (2002) Utilization of l-ascorbate by Escherichia coli K-12: assignments of functions to products of the yjf-sga and yia-sgb operons. J Bacteriol 184, 302–306.
10.1128/JB.184.1.302-306.2002
CASPubMedWeb of Science®Google Scholar
151 Zhang Z, Aboulwafa M, Smith MH & Saier MH Jr (2003) The ascorbate transporter of Escherichia coli. J Bacteriol 185, 2243–2250.
10.1128/JB.185.7.2243-2250.2003
CASPubMedWeb of Science®Google Scholar
152 Campos E, Aguilar J, Baldoma L & Badia J (2002) The gene yjfQ encodes the repressor of the yjfR-X regulon (ula), which is involved in l-ascorbate metabolism in Escherichia coli. J Bacteriol 184, 6065–6068.
10.1128/JB.184.21.6065-6068.2002
CASPubMedGoogle Scholar
153 Campos E, Baldoma L, Aguilar J & Badia J (2004) Regulation of expression of the divergent ulaG and ulaABCDEF operons involved in l-ascorbate dissimilation in Escherichia coli. J Bacteriol 186, 1720–1728.
10.1128/JB.186.6.1720-1728.2004
CASPubMedWeb of Science®Google Scholar
154 Forouhar F, Lee I, Benach J, Kulkarni K, Xiao R, Acton TB, Montelione GT & Tong L (2004) A novel NAD-binding protein revealed by the crystal structure of 2,3-diketo-l-gulonate reductase (YiaK). J Biol Chem 279, 13148–13155.
10.1074/jbc.M313580200
CASPubMedGoogle Scholar

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Volume274, Issue1

January 2007

Pages 1-22

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Vitamin C

Biosynthesis, recycling and degradation in mammals

Abstract

Abbreviations

Formation of vitamin C in mammals and other vertebrates

Outline of the pathway

Effect of xenobiotics on vitamin C formation

Formation of glucuronate from UDP-glucuronate

Glucuronate reductase (aldehyde reductase)

Urono- and gulonolactonase

L-Gulonolactone oxidase

Characterization of the enzyme and of the catalyzed reaction

Molecular defects in man and other species

An alternative fate for glucuronate: the pentose pathway

Control of vitamin C synthesis

Outline on the regulation of vitamin C synthesis

Effect of xenobiotics

Effect of glutathione

Recycling of vitamin C

Reduction of semidehydroascorbate

NADH-cytochrome b5 reductase

Thioredoxin reductase

Cytochrome b561-mediated reduction of intravesicular SDA

Reduction of extracellular SDA involving transfer of electrons across the plasma membrane

Reduction of dehydroascorbate

Spontaneous reaction with GSH

Glutaredoxin and protein disulfide isomerase

Omega class glutathione transferase

3α-Hydroxysteroid dehydrogenase

Thioredoxin reductase

Assessment of the role of the reductive systems

Catabolism of vitamin C

Spontaneous breakdown of dehydroascorbate

Degradation of vitamin C in mammals

In vivo studies

In vitro studies

Utilization of vitamin C by Escherichia coli

Acknowledgements

References

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Figures

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NADH-cytochrome b₅ reductase

Cytochrome b₅₆₁-mediated reduction of intravesicular SDA