Journal of Experimental Botany, Vol. 51, No. 345, pp. 669-674,
April 2000
© 2000 Oxford University Press
The control of ascorbic acid synthesis and turnover in pea seedlings
School of Biological Sciences, University of Exeter, Hatherly Laboratories, Prince of Wales Road, Exeter EX4 4PS, UK
Received 10 September 1999; Accepted 2 December 1999
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Abstract |
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The rate of ascorbate synthesis and turnover in pea seedling embryonic axes was investigated in relation to its pool size. Ascorbate accumulated in embryonic axes of germinating pea seeds which has been supplied with ascorbate. Incorporation of [U-14C]glucose into ascorbate after a 2 h labelling period was reduced by ascorbate loading for 3 h and 20 h, providing evidence that ascorbate biosynthesis is inhibited by endogenous ascorbate. Ascorbate turnover was estimated by following the metabolism of [1-14C]ascorbate over 2 h after ascorbate loading and by the rate of decrease of the ascorbate pool size after ascorbate loading. Ascorbate turnover rate, determined by [1-14C]- ascorbate metabolism, increased as a linear function of pool size. The absolute turnover rate was higher in ascorbate-loaded embryonic axes but was always about 13% of the pool per hour. The initial rate of ascorbate turnover, estimated from the net decrease in pool size after ascorbate loading, also showed a similar turnover rate to that estimated from [1-14C]- ascorbate metabolism. Ascorbate loading had no effect on ascorbate peroxidase, monodehydroascorbate reductase, dehydroascorbate reductase or glutathione reductase activity. Ascorbate oxidase activity decreased after ascorbate loading.
Key words: Antioxidants, ascorbate, biosynthesis, metabolism, vitamin C.
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Introduction |
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L-Ascorbic acid (AsA) is an important component of the plant antioxidant system (Smirnoff, 1996; Noctor and Foyer, 1998; Smirnoff and Wheeler, 1999). Its key role is illustrated by an Arabidopsis thaliana mutant (vtc1) which contains 30% of wild type AsA and is hypersensitive to oxidative stresses caused by ozone and UV-B radiation (Conklin et al., 1996
). In addition to its well-established antioxidant role, AsA has been proposed to have roles in regulation of photosynthesis (Noctor and Foyer, 1998), cell expansion (Smirnoff, 1996) and trans-membrane electron transport (Horemans et al., 1994). It is of interest to understand the biosynthesis of AsA and the factors determining its pool size because of its importance as an antioxidant and also because plants are the major source of dietary vitamin C for humans. The biosynthetic pathway of AsA in plants has been poorly understood until recently (Loewus, 1988; Smirnoff and Wheeler, 1999) and, consequently, very little is known about control of its pool size. A biosynthetic pathway has now been proposed in which AsA is synthesized from L-galactose and L-galactono-1,4-lactone (L-GAL) in a two-step oxidation catalysed by L-galactose dehydrogenase and L-GAL dehydrogenase (Østergaard et al., 1997; Wheeler et al., 1998). L-Galactose is formed from GDP-mannose and GDP-L-galactose (Wheeler et al., 1998). Genetic evidence that mannose is involved in AsA synthesis has been provided by the identification of VTC1 from the AsA-deficient A. thaliana mutant vtc1 as GDP-mannose pyrophosphorylase (Conklin et al., 1997, 1999).
Identification of the biosynthetic pathway provides an opportunity to manipulate the AsA concentration in plants. However, to achieve this goal, it is also necessary to understand the factors controlling AsA synthesis and turnover in various types of tissue. AsA pool size is the result of the balance between the rates of synthesis and turnover. Turnover results from further metabolism or from oxidation followed by non-enzymatic degradation. AsA is readily oxidized to monodehydroascorbate (MDA) as part of its antioxidant function. Oxidation is catalysed by two enzymes: ascorbate peroxidase (APX) and ascorbate oxidase (AO). MDA disproportionates to dehydroascorbate (DHA) and AsA if it is not immediately reduced. DHA is unstable above pH 7 and irreversibly delactonizes to 2,3-diketogulonate (Loewus, 1988; Smirnoff, 1995). Under normal circumstances the AsA pool is at least 90% reduced. This is achieved by the action of two enzymes: NAD(P)-dependent monodehydroascorbate reductase (MDAR) and glutathione (GSH)-dependent DHAR. Along with GR, which regenerates GSH, these enzymes comprise the AsA-GSH cycle (Noctor and Foyer, 1998). High activities of AsA-GSH cycle enzymes could stabilize AsA by preventing DHA accumulation (Foyer et al., 1995). Metabolic products of AsA include oxalate and tartrate but the enzymes involved have not been identified (Loewus, 1988; Saito, 1996). Labelling experiments have shown that AsA is subject to turnover. This has been estimated from 14C-AsA metabolism by leaves of wild-type A. thaliana and the biosynthesis mutant (vtc1) which contains 30% of wild-type AsA concentration. In both cases about 40% of the total pool was metabolized in 22 h. The absolute rate of turnover was higher in wild-type plants, suggesting that AsA turnover is proportional to pool size (Conklin et al., 1997). The AsA pool is larger in leaves grown at high light intensities (Grace and Logan, 1996; Smirnoff and Pallanca, 1996). The AsA concentration decreases in detached A. thaliana and barley leaves during dark periods (Smirnoff and Pallanca, 1996; Conklin et al., 1997) and this is partially reversed by sugar feeding (Smirnoff and Pallanca, 1996). In contrast, the AsA pool of non-photosynthetic tissues is less affected by carbohydrate status because AsA in young embryonic axes of germinating pea seeds is unaffected by carbohydrate feeding or starvation (Pallanca and Smirnoff, 1999).
The results summarized above suggest that AsA pool size is determined both by its rate of synthesis and by its rate of turnover. The capacity of the AsA-GSH cycle and the rate of its catabolism to products such as oxalate and tartrate both contribute to breakdown. The objectives of the experiments described here were to determine the effect of AsA pool size on its rate of synthesis and turnover in non-photosynthetic tissue. Ascorbate metabolism has been characterized in pea seedlings. The seeds contain very little AsA but it accumulates rapidly between 20–40 h after imbibition (Pallanca and Smirnoff, 1999). To date there have been no quantitative estimates of the rates of AsA biosynthesis and turnover or of the effect of AsA pool size on these processes. The rapid synthesis of AsA makes pea embryonic axes suitable for investigating its control. The approach reported here was to increase the AsA pool by loading the tissue with AsA and then to estimate the rates of AsA synthesis and turnover by labelling with 14C-glucose and 14C-AsA. The results provide evidence that AsA synthesis is strongly inhibited by increased AsA pool size while the rate of AsA turnover is increased.
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Materials and methods |
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Plant culture
Pea seeds (Pisum sativum L. cv. Meteor) were surface-sterilized for 10 min in sodium hypochlorite (1.2% available chlorine). After sterilization the seeds were put into plastic boxes between paper towelling moistened with a 20% strength nitrate-type Long Ashton nutrient medium (Hewitt and Smith, 1975
) and germinated in the dark at 21 °C. AsA or L-GAL were added to the nutrient medium at the beginning of imbibition in some experiments. In other cases the seedlings were germinated for 24 h and then separated into cotyledons and embryonic axes. After dissection from seedlings, embryonic axes were incubated in glass vials on two layers of 1 cm diameter filter paper discs moistened with 0.4 ml sterile nutrient medium with the desired additions. The vials were enclosed in plastic boxes and incubated in the dark at 21 °C.
Ascorbate assay
Embryonic axes (approximately 12–14 per sample) were frozen in liquid nitrogen, ground to a powder and extracted in 1 ml 5% perchloric acid. The homogenate was centrifuged at 12 000 g for 2 min and the supernatant was neutralized with 5 M potassium carbonate using methyl orange as indicator. The neutralized supernatant was centrifuged again and used for AsA assay with AO (Hewitt and Dickes, 1961). Total AsA was assayed after reducing the sample with homocysteine (Okamura, 1980). DHA concentration was calculated from the difference between total AsA and AsA.
Metabolism of 14C-labelled glucose and ascorbate by embryonic axes
D-[U-14C]Glucose (111 kBq, specific activity 10.9 GBq mmol-1, Amersham) and L-[1-14C]AsA (3.7 kBq, specific activity 18.5 MBq mmol-1, Amersham) were fed to embryonic axes as described above. The labelled glucose, if all absorbed, would have increased the glucose concentration in the embryonic axes by 0.05 µmol g-1 fresh weight. After incubation the samples (each containing 12–14 embryonic axes) were extracted in perchloric acid as described above and the supernatant was separated into neutral, acidic and basic fractions by ion exchange. AsA was isolated in a fraction eluted from a strong anion exchange resin by 60 mM formic acid. It was then purified by HPLC and its 14C content determined with a liquid scintillation counter (Conklin et al., 1997).
Enzyme assays
MDAR and DHAR were extracted by homogenizing 10 embryonic axes in 1 ml of the medium described by Smirnoff and Colombé (Smirnoff and Colombé, 1988). APX and AO were extracted as described by Conklin et al. (Conklin et al., 1997). MDAR and APX were assayed by the method of Smirnoff and Colombé (Smirnoff and Colombé, 1988) and DHAR by the method of Nakano and Asada (Nakano and Asada, 1980). An uncentrifuged tissue homogenate was used to measure soluble and cell wall-bound AO by the method of Conklin et al. (Conklin et al., 1997). Protein was measured by the Coomassie blue binding method (Bradford, 1976).
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Results |
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Ascorbate synthesis from [U-14C]glucose in relation to pool size
To determine the effect of AsA pool size on the rate of AsA synthesis from glucose, the seedlings or embryonic axes were loaded for either 3 h or 20 h with 50 mM AsA. These treatments increased AsA pool size to 4.3 and 6.3 µmol g-1 fresh weight, respectively (Table 1
). The oxidation state of the AsA pool was not affected by AsA loading. In all cases DHA was about 10% of the total pool, therefore only total AsA concentration is shown. After ascorbate loading the tissues were labelled with [U-14C]glucose for 2 h and the incorporation of 14C into AsA was determined. AsA loading had no effect on [U-14C]glucose uptake (Table 1). However, AsA loading decreased the percentage of [U-14C]glucose incorporated into AsA by 59% after 3 h loading and by 83% after 20 h loading (Table 1). There were small but statistically significant effects of ascorbate loading on the distribution of 14C in the neutral+basic, acidic, insoluble, and CO2 fractions (Table 1). The relative rate of labelling of AsA was linearly related to tissue AsA concentration (Fig. 1) even though the length of exposure to AsA differed between treatments. The results show that the apparent rate of AsA synthesis is decreased by high AsA and that the effect is appreciable even after a short loading period.
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Ascorbate turnover in relation to pool size
AsA turnover was estimated by measuring the breakdown of [1-14C]AsA and by the rate of net loss of AsA from embryonic axes loaded with AsA. The seedlings or embryonic axes were first loaded for either 3 h or 20 h with 50 mM AsA. These treatments increased AsA pool size to 3.5 and 6.2 µmol g-1 fresh weight, respectively (Table 2
). The loaded tissue was then transferred to [1-14C]AsA for 2 h to follow metabolism of labelled AsA. 14C-AsA uptake was significantly reduced by 30% after AsA loading. 40–50% of the total 14C remained in AsA after 2 h. Most of the label was distributed between the acidic and neutral+basic fractions. Only 2% of the 14C was released as CO2. This pattern of labelling was not significantly affected by AsA loading (Table 2). The same proportion (40–50%) of 14C-AsA was metabolized in tissues containing high or low AsA pool sizes (Table 2). The amount of 14C appearing in compounds other than AsA was used to calculate AsA turnover after correcting for the mean specific activity of AsA in each treatment (Conklin et al., 1997). This showed that the turnover rate is a linear function of pool size, irrespective of the AsA loading time (Fig. 1). The mean turnover rate of AsA as a percentage of the total pool size over all AsA concentrations was 13.1±2.7% h-1.
To assess the validity of the results from 14C-AsA metabolism, the AsA content of embryonic axes loaded for 24 h with 50 mM AsA was followed after transferring the seedlings to AsA-free medium. The AsA content of the embryonic axes decreased after transfer to AsA-free medium (Fig. 2). The rate of loss calculated from the [1-14C]AsA labelling experiment (13% h-1) predicts a rate of loss very similar to that observed, particularly in the first few hours (Fig. 2). After 3–4 h the rate of loss of AsA deviated slightly from that predicted by the [1-14C]AsA labelling experiment. This is explained by the progressive dilution of the AsA concentration by growth of the embryonic axes over the 6 h period (content per embryonic axis and not concentration is shown in Fig. 2). After 24 h the AsA content of AsA-loaded embryonic axes (0.58±0.08 nmol embryonic axis-1) was still higher than control embryonic axes (0.12±0.03 nmol embryonic axis-1).
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). Values are means±standard deviation (n=3).
The effect of ascorbate loading on the activity of ascorbate metabolizing enzymes
The increased rate of AsA breakdown in AsA-loaded tissue could be caused by changes in activity of enzymes which catalyse AsA oxidation (APX and AO) and ascorbate-glutathione cycle enzymes (MDAR, DHAR and GR) which regenerate AsA from its oxidized forms. APX, MDAR, DHAR, and GR did not change in activity after AsA-loading (Table 3). In contrast, AO activity decreased markedly after AsA loading. After imbibing pea seedlings with AsA or L-GAL, the immediate biosynthetic precursor of AsA (Smirnoff and Wheeler, 1999), for 24 h the activity of AO in the embryonic axes decreased. The reduction in activity correlated with AsA increasing concentration in the embryonic axes (Fig. 3), however, further investigation is required to determine the cause of this striking effect.
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Discussion |
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The control of AsA metabolism has not been investigated in much detail so far largely because the biosynthetic pathway has only recently been established (Wheeler et al, 1998
; Conklin et al., 1999). The aim of this investigation was to establish if AsA biosynthesis and turnover are influenced by the endogenous AsA pool by using labelled substrates to estimate the rate of AsA synthesis and turnover. The results show that AsA synthesis is strongly inhibited by a large AsA pool, while turnover rate is faster and directly proportional to pool size.
Control of ascorbate biosynthesis
The rate of AsA synthesis from 14C-glucose was decreased after elevating the pool size by exogenous AsA. This conclusion still remains true after correcting for the proportion of AsA metabolized over this period, which was estimated from 14C-AsA metabolism, and is similar (13% h-1) in all treatments. The rate of AsA synthesis decreases as a linear function of pool size, irrespective of time of exposure to exogenous AsA. Because inhibition of synthesis was detected within 3 h of AsA feeding, it could be caused by direct feedback inhibition of AsA biosynthesis enzymes. Further information is required to distinguish between this possibility and repression of the expression of biosynthesis enzymes. This is the first quantitative demonstration of ‘feedback’ control of AsA biosynthesis. It has been shown that exogenous glucose was more effective at elevating AsA concentration in potato tubers containing initially low AsA than those containing high AsA (De Gara et al., 1989), an observation which provides indirect evidence for ‘feedback’ control. Now that the biosynthetic enzymes have been identified (Wheeler et al., 1998; Conklin et al., 1999) it will be possible to study their regulatory properties and expression and, therefore, to identify which steps are subject to feedback inhibition or repression.
Ascorbate turnover
AsA turnover rate was estimated by metabolism of [1-14C]AsA and by following the net loss of AsA from AsA-loaded embryonic axes. Considering the uncertainties that are inherent in the estimation of AsA turnover, it is noteworthy that both methods gave similar results, particularly over the first 2 h of measurement, and showed that the absolute rate of turnover is directly proportional to pool size. The uncertainties include oxidation and non-enzymatic loss of DHA and the possibility that the distribution of the exogenous AsA between subcellular compartments is not the same as the endogenous pool. This could result in different susceptibility of exogenous and endogenous pools to oxidation and breakdown. However, the conclusion that the measurements provide a reliable estimate of AsA turnover is supported by the application of the same techniques to A. thaliana leaves. In this case different pool sizes could be compared without AsA loading by using the vtc1 mutant which has 30% of the wild-type AsA content (Conklin et al., 1997). AsA turnover was estimated by [1-14C]AsA metabolism and the rate of decrease of the pool in the dark. Both methods gave the same result and, as with pea embryonic axes, showed that the rate of breakdown is proportional to pool size. The estimated turnover rate in embryonic axes with a normal pool size is 0.2 µmol h-1 g-1 fresh weight, representing about 13% of the pool per hour. AsA turnover in wild-type A. thaliana is 0.1 µmol h-1 g-1 fresh weight at low light intensity, representing about 2.5% of the pool per hour (Conklin et al., 1997). It would be expected that the pattern of AsA metabolism differs between leaves and non-photosynthetic tissues. For example, the ascorbate pool of leaves is increased by sugar supply in the dark (Smirnoff and Pallanca, 1996), while the AsA pool of embryonic axes is not affected (Pallanca and Smirnoff, 1999). The higher turnover in the embryonic axes may reflect their faster growth rate.
AsA turnover could result from non-enzymatic DHA breakdown or from metabolism of AsA or DHA to oxalate and tartrate (Loewus, 1988). A model in which increased DHA concentration results in increased turnover could be proposed since DHA is unstable above pH 7. The AsA loading experiments showed that the embryonic axes have a sufficiently high capacity to maintain the AsA pool at least 90% reduced, however, the absolute DHA concentration increases on AsA loading. DHA breakdown would therefore follow the same first order kinetics observed in the turnover measurements. The importance of DHA breakdown in causing AsA turnover is supported by the increased leaf AsA pool found in transgenic poplar over-expressing GR (Foyer et al., 1995). APX and AO activities were not induced by AsA loading. On the contrary, AO activity was strongly reduced. Therefore increased turnover rate cannot be attributed to higher capacities of these enzymes. There is currently no explanation for the strong decrease in AO activity in AsA-loaded tissue and it would be premature to speculate. Enzymatic conversion of AsA to other metabolic products such as oxalate and tartrate will also contribute to AsA turnover. Currently, nothing is known about the enzymes involved in these pathways (Loewus, 1988). Clearly, full understanding of AsA turnover will require more detailed characterization of the products of AsA and DHA metabolism.
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Conclusions |
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AsA pool size is potentially controlled by ‘feedback’ inhibition of synthesis and by turnover. These processes now need to be investigated in more detail. This will provide the information needed to optimize attempts to increase AsA concentration by metabolic engineering in transgenic plants.
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Acknowledgments |
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This research was supported by a BBSRC (Biochemistry of Metabolic Regulation in Plants) grant. We thank Marjorie Raymond for excellent technical support.
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Notes |
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1 To whom correspondence should be addressed. Fax: +44 1392 263700. E-mail:N.Smirnoff{at}exeter.ac.uk
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