JXB Advance Access originally published online on January 13, 2007
Journal of Experimental Botany 2007 58(5):1109-1118; doi:10.1093/jxb/erl269
| ||||||||||||||||||||||||||||||||||||||||||||||||||||||||||||||||||||||||||||||||||||
© 2007 The Author(s).
This is an Open Access article distributed under the terms of the Creative Commons Attribution Non-Commercial License (http://creativecommons.org/licenses/by-nc/2.0/uk/) which permits unrestricted non-commercial use, distribution, and reproduction in any medium, provided the original work is properly cited.This paper is available online free of all access charges (see http://jxb.oxfordjournals.org/open_access.html for further details)
RESEARCH PAPER |
The effect of Glc6P uptake and its subsequent oxidation within pea root plastids on nitrite reduction and glutamate synthesis
1Faculty of Life Sciences, The University of Manchester, 3.614 Stopford Building, Oxford Road, Manchester M13 9PT, UK
2Institute for Protein Research, Osaka University, Osaka 565 0871, Japan
3University of Bristol, Long Ashton Research Station, Bristol BS41 9AF, UK
4University of Groningen, Laboratory of Plant Physiology, PO Box 14, 97500 AA, Haren, The Netherlands
5College of Biological Science, University of Guelph, Guelph N1G 2W1, Canada
* To whom correspondence should be addressed. E-mail: caroline.bowsher{at}manchester.ac.uk
Received 25 July 2006; Revised 14 November 2006 Accepted 16 November 2006
 |
Abstract |
|---|
 Top Abstract IntroductionMaterials and methodsResultsDiscussionReferences |
|---|
In roots, nitrate assimilation is dependent upon a supply of reductant that is initially generated by oxidative metabolism including the pentose phosphate pathway (OPPP). The uptake of nitrite into the plastids and its subsequent reduction by nitrite reductase (NiR) and glutamate synthase (GOGAT) are potentially important control points that may affect nitrate assimilation. To support the operation of the OPPP there is a need for glucose 6-phosphate (Glc6P) to be imported into the plastids by the glucose phosphate translocator (GPT). Competitive inhibitors of Glc6P uptake had little impact on the rate of Glc6P-dependent nitrite reduction. Nitrite uptake into plastids, using 13N labelled nitrite, was shown to be by passive diffusion. Flux through the OPPP during nitrite reduction and glutamate synthesis in purified plastids was followed by monitoring the release of 14CO2 from [1-14C]-Glc6P. The results suggest that the flux through the OPPP is maximal when NiR operates at maximal capacity and could not respond further to the increased demand for reductant caused by the concurrent operation of NiR and GOGAT. Simultaneous nitrite reduction and glutamate synthesis resulted in decreased rates of both enzymatic reactions. The enzyme activity of glucose 6-phosphate dehydrogenase (G6PDH), the enzyme supporting the first step of the OPPP, was induced by external nitrate supply. The maximum catalytic activity of G6PDH was determined to be more than sufficient to support the reductant requirements of both NiR and GOGAT. These data are discussed in terms of competition between NiR and GOGAT for the provision of reductant generated by the OPPP.
Key words: Glucose-6-phosphate dehydrogenase, glucose-6-phosphate uptake, glutamate synthase, nitrite uptake, nitrite reductase maximum catalytic activity, oxidative pentose phosphate pathway competition
|
Introduction |
|---|
|
TopAbstractIntroductionMaterials and methodsResultsDiscussionReferences |
|---|
In higher plants, nitrogen assimilation involves the enzymes nitrate reductase (NR, EC 1.6.6.2 [EC] ), nitrite reductase (NiR, 1.7.7.1 [EC] ), glutamine synthetase (GS, 6.1.1.3 [EC] ), and glutamate synthase (GOGAT, EC 1.4.1.13 [EC] and 1.4.7.1 [EC] ; Crawford, 1995; Lam et al., 1996). There is some divergence in higher plants between those which are considered to be predominantly leaf assimilators, and those which assimilate nitrate mainly in the root such as legumes (Andrews, 1986; Pate, 1980). Nitrate assimilation is a highly compartmentalized system as NR is located in the cytosol, whilst NiR, a proportion of the GS, and all the GOGAT are found in the plastids. The available data from studies on transgenic and genetically manipulated plants indicates that the levels of these enzymes do not limit primary nitrate assimilation and therefore crop yield (Andrews et al., 2004; Good et al., 2004). The process in plastids is also dependent upon a supply of reducing power. Reductant is generated photosynthetically in leaf tissue. Clearly, in non-photosynthetic tissue this is not the case and, indeed, the generation of reducing power by oxidative metabolism could also have an impact on nitrogen assimilation.
There is good evidence that, in root plastids, the pentose phosphate pathway generates reductant in the form of NADPH, which is used subsequently to drive nitrogen assimilatory reactions (Bowsher et al., 1989, 1992). The oxidative reactions of the pentose phosphate pathway (OPPP) are capable of operating in both the plastids and the cytosol (Debnam and Emes, 2000; Schnarrenberger et al., 1995). Whilst there appear to be species differences in the localization of the non-oxidative reactions in the cytosol, the complete pathway has been found in plastids of all species examined thus far. It is the initial conversion of glucose 6-phosphate (Glc6P) to ribose 5-phosphate, catalysed by glucose 6-phosphate dehydrogenase (G6PDH, EC 1.1.1.49 [EC] ) and 6 phosphogluconate dehydrogenase (6PGDH, EC 1.1.1.44 [EC] ), which produces two molecules of NADPH. Six members of the G6PDH gene family have been identified in Arabidopsis thaliana, based on transit peptide analysis it is predicted that there are two cytosolic and four plastidic isoforms (Kruger and von Schaewen, 2003; Wakao and Benning, 2005). Five members of the 6PGDH gene family have also been identified of which at least two of these are thought to be plastidic in location (Kruger and von Schaewen, 2003). Based on the expression of genes encoding G6PDH and 6PGDH from A. thaliana following transfer to nitrate-containing medium, their involvement in providing NADPH for ferredoxin-dependent reactions has been implicated (Wang et al., 2000). A 12–27-fold increase in both enzymes activities in maize root plastids after 24 h treatment with 10 mM nitrate has also been reported (Redinbaugh and Campbell, 1998). Furthermore, increase in the expression of the P2 form of G6PDH from roots of Nicotiana tabacum (Knight et al., 2001) and Lycoperscion esculentum (Wang et al., 2001) in response to nitrate and from Hordeum vulgare root plastids in response to ammonium/glutamate (Esposito et al., 2001b) have also been reported.
The ability to monitor the flux through the OPPP using purified root plastids has allowed the demonstration of increased carbohydrate oxidation in response to nitrite reduction (Bowsher et al., 1989) and glutamate synthesis (Bowsher et al., 1992; Esposito et al., 2003). This approach has also provided information on the stoichiometry of the interaction between pathways of carbon and nitrogen metabolism (Hartwell et al., 1996). Furthermore the P2 form of G6PDH has been confirmed as supporting the increased flux through the OPPP (Esposito et al., 2003). Finally, when Glc6P is added alone or simultaneously with nitrite to barley root plastids, the increased NADPH/NADP ratio observed is consistent with the activation of G6PDH enzyme activity and the OPPP (Wright et al., 1997). The reactions catalysed by NiR and GOGAT both require reduced ferredoxin as a substrate. The OPPP-generated NADPH acts as the initial reductant to generate reduced ferredoxin via a root-specific ferredoxin-NADP oxidoreductase (FNR, EC 1.18.12). FNR transcript and enzyme activity is co-induced with root-specific ferredoxin during the induction of nitrate assimilation (Bowsher et al., 1993; Matsumura et al., 1997; Ritchie et al., 1994).
Using pea root plastids, it has previously been shown that Glc6P can be taken up and supports nitrite reduction and the biosynthesis of glutamate (Bowsher et al., 1989, 1992). In contrast to photosynthetic tissues, there is a need to import carbon into the plastids of non-photosynthetic tissue as a source of energy to drive biosynthetic reactions. Depending on the tissue examined and the plastid type, a range of phosphate translocators has been identified which are capable of importing carbon precursors. The triose phosphate/phosphate translocator (TPT) mediates the export of fixed carbon in the form of triose phosphates and 3-phosphoglycerate in exchange for inorganic phosphate (Fliege et al., 1978). Although it is not clear whether the TPT is expressed in roots (Flugge, 1999; Knight and Gray, 1994), more recently, two novel plastid transporters have been identified which show higher expression in roots. The plastidic PEP/phosphate translocator (PPT), which transports phosphoenolpyruvate (PEP) in exchange with inorganic phosphate (Fischer et al., 1997), and the glucose-6-phosphate/phosphate translocator (GPT), which counter exchanges Glc6P with inorganic phosphate and triose phosphates. This latter carrier is analogous to the chloroplast TPT, but has a much higher affinity for Glc6P (Borchert et al., 1989, Flugge, 1999; Kammerer et al., 1998). At the protein level, the GPT has low but significant similarity to the TPT (Flugge, 1999) and the PPT (Fischer et al., 1997). The proposed physiological function of the GPT is to facilitate the import of Glc6P for use as an OPPP substrate. In the Arabidopsis genome two paralogous GPT genes have been identified, AtGPT1 and AtGPT2 (Knappe et al., 2003). Although both GPTs appear functional in planta, the ubiquitously expressed AtGPT1, like pea GPT, is expressed in roots and is thought to be the major GPT involved in Glc6P transport into heterotrophic plastids. Loss of AtGPT1 function disrupts the OPPP and has been proposed to affect the reducing power supply needed for fatty acid biosynthesis, leading to less lipid body formation and disintegration of membrane systems, resulting in impaired pollen maturation and embryo sac development (Niewiadomski et al., 2005). Glc6P import into plastids potentially impacts not only the capacity of the plastids to sustain the OPPP, but also other related metabolic processes including nitrate assimilation via NiR and GOGAT.
The reduction of nitrite and synthesis of glutamate are dependent upon both the uptake of Glc6P and its subsequent oxidation within the organelle. In this paper, the relative importance of these two aspects in controlling nitrogen assimilation using purified root plastids of Pisum sativum is investigated. The approaches taken in answering these questions were (i) to determine the effect of inhibition of Glc6P uptake on nitrite reduction; (ii) to investigate the mechanism of nitrite entry into the organelle; and (iii) to determine whether there is competition for reductant between the processes of nitrite reduction and glutamate synthesis. The results demonstrate that there is competition for electrons between nitrite reductase and glutamate synthase and that the OPPP is not able to produce sufficient reductant to support the optimal activities of these two enzymes simultaneously. This suggests that the oxidation of Glc6P through the OPPP may place a significant constraint on the reactions of nitrogen assimilation in plastids.
|
Materials and methods |
|---|
|
TopAbstractIntroductionMaterials and methodsResultsDiscussionReferences |
|---|
Chemicals
All reagents were of ‘AR’ grade where possible. Cofactors, enzymes and substrates were purchased from BDH, Poole, Dorset (UK), Boehringer Mannheim (FRG), or Sigma, Poole, Dorset, (UK). Percoll was purchased from Amersham Pharmacia, Milton Keynes (UK). [1-14C]-glucose-6-phosphate was from New England Nuclear, Southampton (UK) and Silicone AR200 oil was from Water Chemicals UK Ltd.
Plant material
Pea (Pisum sativum L. cv. Early onward) seeds purchased from Yates Seed Breeders and Merchants (Macclesfield, Cheshire UK), were germinated in the dark for 5 d at 25 °C as described previously (Emes and England, 1986). On day 5, 24 h prior to extraction, the peas were watered three times with 50 mM KNO3.
Extraction and purification of root plastids
Plastids were prepared from 150–300 g of roots, based on the method described previously (Bowsher et al., 1989). Intact plastids were maintained in buffer A containing 50 mM N-[2-hydroxy-1,1-bis(hydroxymethyl)ethyl]glycine (Tricine-NaOH, pH 7.9, containing 330 mM sorbitol, 1 mM ethyldiaminetetra-acetic acid (EDTA), 1 mM MgCl2). When required, ‘broken’ plastids were ruptured osmotically by resuspending in distilled water, followed by the addition of an equal volume of double-strength buffer A to standardize the salt concentration. In some instances Triton X-100 was added to a final concentration of 0.1% (v/v), although this had no effect on the results obtained.
Glucose-6-phosphate-dependent nitrite reductase/glutamate synthase activity
The assay contained 150 mM 2-amino-2-(hydroxymethyl)-1,3-propanediol (Tris)-HCl, pH 8.0, 0–10 mM glucose-6-phosphate, substrate (1 mM nitrite, for nitrite reductase and/or 5 mM glutamine and 5 mM 2-oxoglutarate for glutamate synthase) and intact or broken plastids (550 µg protein). All solutions were osmotically buffered with 330 mM sorbitol. At known times the reaction was stopped by the addition of either 0.5% (v/v) acetic acid or 0.4 mM 5-sulphosalicylic acid, respectively. Samples were removed for nitrite determination or glutamate estimation as described previously (Bowsher et al., 1989, 1992). All assays were corrected for nitrite reduction or glutamate synthesis in broken plastids by carrying out the same experiments using organelles osmotically lysed and subtracting these values from those obtained with intact preparations.
Glucose-6-phosphate dehydrogenase
G6PDH was assayed based on the method of Fowler and ap Rees (1970).
Protein determination
Protein was measured using a commercial Bio-Rad (Watford, UK) protein method based on Bradford (1976) with thyroglobulin as the protein standard.
Measurement of 14CO2 evolution from [1-14C]-glucose-6-phosphate
All assays were carried out in a final volume of 1 cm3 in the presence of nitrite and/or glutamine and 2-oxoglutarate as previously described (Bowsher et al., 1989, 1992).
Transport measurements
Silicone oil centrifugation was carried out based on the method of Emes and Traska (1987). All reagents were made up in 330 mM sorbitol. For Glc6P uptake measurements, 90 µl root plastids (approximately 100 µg protein, resuspended in buffer A) were incubated for 15 min at room temperature with 10 µl 330 mM sorbitol, or 10 µl 330 mM sorbitol containing DIDS, phosphoglyceric acid (PGA), inorganic orthophosphate (Pi), or dihydroxyacetone phosphate (DHAP). At the end of this period, U-[14C]-Glc6P (specific activity=3.7 kBq µmol–1) was added to the mix for 30 s at 20 °C. Substrate uptake was stopped by centrifugation at 13 000 g for 120 s through 50 µl silicone oil (AR200). Pelleted material was removed by excising the bottom of the tube. The lower portion of the cut tube was placed in Ecoscint scintillation fluid and thoroughly shaken before radioisotope counting. Residual silicone oil did not cause an increase in quenching. The volume of organelles sedimented was calculated from measurements with tritiated water and corrected for the organelle-free space with 14C-dextran (Heldt, 1980). This latter value varied between 2% and 6% of the space occupied by 3H2O in the sediment. Estimation of the ‘sorbitol-permeable’ space with U-14C sorbitol gave values which were not significantly different from those obtained with 14C-dextran.
To determine nitrite uptake a 6.5 ml aliquot of 13N-labelled nitrate (prepared by proton irradiation of pure water at the Positron Emission Tomography Centre, University Hospital, Groningen, The Netherlands) was first converted to 13N-nitrite based on the method developed by Brewer et al. (1965) in which nitrate is reduced in the presence of cadmium and EDTA (see Grasshoff, 1964; Wood et al., 1967, for a discussion of the chemistry). One ml of nitrate/EDTA solution (2.80g Na2-EDTA, 246 µl 1000 ppm KNO3, adjusted to pH 6.8 with KOH and made up to 25 ml) was pipetted into a 10 ml glass vial in a lead pot. A 6.5 ml aliquot of 13N-labelled nitrate, that also contained some 13
(prepared by proton irradiation of pure water at the Positron Emission Tomography Centre, University Hospital, Groningen, The Netherlands), was withdrawn via a hypodermic needle from the sealed multidose glass bottle, located within a lead pot, into a 25 ml syringe, which itself was housed in a lead jacket with lead glass window and added, with mixing, to the vial to give a solution of 100 µmol KNO3 in 1.5% (w/v) EDTA.
The solution containing 13
, 13
, unlabelled , and EDTA; was pumped by a peristaltic pump at 0.32 ml min–1 through a cadmium reduction coil. The coil consisted of an 80 cm length of polythene tubing, internal diameter 2.45 mm (Portex Ltd., Hythe, Kent, size 800/500/540) filled with cadmium filings (produced by rasp from stick cadmium metal and gently sieved to remove the smaller particles that would pass through a size 35 mesh). The coil was lightly plugged at each end with silica wool and kept filled with either water or the solution from the previous run to prevent oxidation and to maintain chemical reactivity. At the start of the reduction process the water or previous solution entrapped in the coil was flushed to waste by passing the fresh 13N-labelled mixture through for about 7 min (until the outflow was strongly radioactive). A resin cation exchange column (1 ml zeo-karb 225, in a 1 ml disposable syringe body) was then rinsed for 1–2 min by the following aliquot of solution. The subsequent 2–2.5 ml of the solution was collected in a polythene scintillation vial insert in a double lead transport container, in which it was conveyed to the experimental location. The final 2–3 ml of the treated solution was collected and frozen for later analysis of nitrite and cadmium concentrations.
Samples were retained for analysis of nitrite and stored frozen for 1–2 d. After thawing, a 0.25 ml aliquot of the sample was diluted to 1 ml with water and nitrite determined using diazo-dye coupling reagents as described previously (Bowsher et al., 1989).
For the determination of cadmium content, the sample was diluted 1:100 or 1:50 and the atomic absorption was measured at 228.8 nm (on a Fisons Atomic Absorption Spectrophotometer, at the Institute of Soil Fertility, Groningen) and compared with a 0–3 µg Cd2+ ml–1 Standard (0.267 µmol).
The uptake of 13N-nitrite into plastids was then determined by incubating root plastids with 0–125 µM 13N-labelled nitrite for 30 s at 20 °C and subsequent determinations made as described above, accounting for the number of half-lives of 13N lost during the procedure.
|
Results |
|---|
|
TopAbstractIntroductionMaterials and methodsResultsDiscussionReferences |
|---|
It has previously been demonstrated that exogenous glucose-6-phosphate (Glc6P) is capable of supporting nitrite reduction and glutamate formation in pea root plastids (Bowsher et al., 1989, 1992; Emes and Fowler, 1983). The importance of the Glc6P/phosphate translocator (GTP) in controlling the Glc6P-dependent nitrite reduction was examined in purified plastids by using substrates which are competitive biochemical inhibitors of the transporter (Figs 1, 2). As would be expected of a protein with phosphate binding activity, DIDS was a strong inhibitor of Glc6P uptake, and hence nitrite reduction (Fig. 1). Because the effect of DIDS is time-dependent and involves covalent modification of the target protein, it was not possible to compare directly the effects of DIDS on Glc6P uptake (which is measured over seconds) with its effect on Glc6P-dependent nitrite reduction (measured over tens of minutes). Consequently, the choice was made to use known physiological competitive inhibitors of Glc6P uptake in order to determine their relative impact on nitrite reduction and Glc6P uptake. Figure 2 shows that Glc6P transport is inhibited by PGA, Pi, and dihydroxyacetone phosphate, producing approximately 50% inhibition at the highest concentrations used. By contrast, none of these compounds had any significant effect on the rate of Glc6P-dependent nitrite reduction, even at concentrations up to 20 mM (Fig. 3a, b, c). The implication of these results is that a substantial decrease in the capacity for Glc6P uptake has little impact on Glc6P-coupled nitrite reduction which is being measured within the organelle. It would seem, therefore, that the GPT exerts little overall control over the process of nitrite reduction. Likewise there was no effect by any of these compounds on 14CO2 evolution from [l-14C]-Glc6P (diagnostic of the flux through the OPPP; Bowsher et al., 1989) supporting the view that the activity of the OPPP was unimpaired by the decrease in the capacity for Glc6P uptake which these competitive inhibitors would produce (Fig. 3 d, e, f).
|
|
|
A second site of possible regulation of Glc6P-dependent nitrite reduction is the movement of nitrite across the plastid envelope. Uptake of
, however, appeared to be linear with concentration and did not saturate at concentrations likely to be encountered in vivo even in excess of 100 µM (Fig. 4). The data suggest that crosses the plastid envelope by passive diffusion. Certainly, there was no direct evidence for a carrier-mediated process and uptake was not impaired in the presence of mM concentrations of other anions such as Pi and 
, nor by DIDS (data not shown). Furthermore, when 13 uptake was measured in the presence of 10 mM Glc6P a more than 3-fold increase in the rate of diffusion of nitrite into the plastids was recorded (data not shown). This suggests that an increase in the availability of Glc6P to support reduction was increasing the diffusion gradient of across the plastid envelope.
|
The above results suggest that the process of nitrite reduction is most likely limited by events within the organelle, rather than by the entry of substrates. Using isotopically labelled Glc6P it is possible to investigate the effect of nitrite reduction and/or glutamate synthesis on carbohydrate oxidation by measuring the release of 14CO2. Previously, it has been shown that nitrite reduction and glutamate synthesis in root plastids are directly responsible for an increase in CO2 evolution specifically and only from C atom 1 of Glc6P, implying an increased flux of C through the OPPP (Bowsher et al., 1989, 1992). The relationship between flux through the OPPP, as measured by 14CO2 evolution from [1-14C]-Glc6P (Fig. 5a), and the ability of the plastid preparations to support reductive assimilation and biosynthesis measured as nitrite reduction (Fig. 5b) and glutamate formation (Fig. 5c) were therefore examined. Figure 5a demonstrates that the evolution of 1-14CO2 increases with Glc6P concentration in the presence of the NiR substrate, nitrite, or in the presence of the GOGAT substrates, glutamine and 2-oxoglutarate. Both the NiR and GOGAT reactions are dependent on a supply of reduced ferredoxin which is initiated by the generation of NADPH through the OPPP. The response to varied Glc6P presumably reflects, at least in part, the kinetics of G6PDH and the other enzymes of the OPPP in utilizing the substrate and products formed as has been observed previously (Bowsher et al., 1989). Evolution of 1-14CO2 was less in the presence of glutamine and 2-oxoglutarate when compared with nitrite, presumably reflecting the relative maximal catalytic activities of NiR and GOGAT. The reduction of 1 mol of nitrite requires six electrons, whereas the conversion of glutamine and 2-oxoglutarate to 2 mol of glutamate requires only two electrons. To investigate whether there is competition between NiR and GOGAT for reductant when operating simultaneously, the impact of both reactions on flux through the OPPP was examined. When plastids were supplied with saturating substrate concentrations for NiR, (nitrite), and GOGAT, (glutamine and 2-oxoglutarate), then at all concentrations of Glc6P, the amount of CO2 released was only equivalent to the rate seen when nitrite reduction alone was occurring (Fig. 5a). These results suggest that at any given operating concentration of Glc6P, irrespective of the demand for reductant, the flux through the OPPP cannot be increased above that which is generated when supporting nitrite reduction alone, even when there is an additional ‘sink’ for electrons effected by the substrates for GOGAT. Experiments were therefore undertaken to measure the impact glutamate synthesis has on Glc6P-dependent nitrite reduction and vice versa. In Fig. 5b it can be seen that the rate of nitrite reduction is decreased in the presence of the substrates for the GOGAT reaction, glutamine and 2-oxoglutarate. This was true at all concentrations of Glc6P, including substrate levels well in excess of the Km for Glc6P uptake or G6PDH. Similarly, at all concentrations of Glc6P, the formation of glutamate was reduced by incubating plastids with the substrate for nitrite reduction in addition to glutamine and 2-oxoglutarate (Fig. 5c).
|
It has previously been shown that the increase in GOGAT activity that occurs through supplying plastids with an increasing glutamine concentration in the presence of a fixed amount of 2-oxoglutarate, leads to an increased flux of carbon through the OPPP (Bowsher et al., 1992). This method of titrating in GOGAT activity by increasing the availability of glutamine substrate was exploited to examine the competition between NiR and GOGAT for reducing power generated by the OPPP (Fig. 6). Intact plastids were supplied with 1 mM nitrite and nitrite reduction and the OPPP flux were followed at high (5 mM) and low (0.5 mM) concentrations of Glc6P. The activity of GOGAT increased with the concentration of glutamine against a fixed amount of 2-oxoglutarate at both Glc6P concentrations (Fig. 6a). Figure 6b shows that the evolution of 14CO2 from 1-14C-Glc6P, and hence the OPPP flux, was unaffected by increasing the glutamine level. By contrast, it was clear that there was a steady decrease in nitrite reduction as the glutamine concentration was raised at either Glc6P concentration used (Fig. 6c). In effect, the GOGAT reaction was being titrated against nitrite reduction in a competition for reducing power.
|
The results of these experiments showed that there was competition for electrons between nitrite reductase and GOGAT, implying that the OPPP is not producing sufficient reductant to support the optimal activities of these two enzymes simultaneously. One way in which the supply of reductant may be restricted is by the activity of enzymes of the OPPP. Previously the activity of 6PGDH in pea root plastids has been shown to increase significantly when roots are incubated in nitrate prior to harvesting (Emes and Fowler, 1983). Glucose-6-phosphate dehydrogenase (G6PDH) is also part constitutive and part induced and it seems that the activity of G6PDH is closely linked to the activities of the inducible nitrogen assimilatory enzymes (Emes and Fowler, 1979; Redinbaugh and Campbell, 1998). G6PDH activity in plastids isolated from pea roots grown in the absence of nitrate was 2556 nmol mg–1 protein h–1. This increased with the nitrate concentration supplied to intact roots 24 h prior to harvesting, reaching a maximum activity of 8150 nmol mg–1 protein h–1 between 10–25 mM external nitrate. The co-ordinated induction of G6PDH activity is clearly a response to the increased demand for reducing power as a result of the induction of nitrate assimilation.
|
Discussion |
|---|
|
TopAbstractIntroductionMaterials and methodsResultsDiscussionReferences |
|---|
In roots, nitrate assimilation is dependent upon a supply of reductant initially generated by oxidative metabolism, a significant portion of which is generated by the OPPP. Nitrate is reduced to nitrite in the cytosol. The uptake of nitrite into the plastids and its subsequent reduction by nitrite reductase and glutamate synthase are potentially important control points. The reduction of nitrite and synthesis of glutamate within the plastid are dependent upon both the uptake of Glc6P and its subsequent oxidation. As such, either of these processes could limit the capacity for nitrogen assimilation. In this paper the relative importance of Glc6P transport and the OPPP in controlling nitrogen assimilatory enzymes dependent on oxidative metabolism in root plastids has been examined.
Pea root plastids are able to transport glucose-6-phosphate in counter exchange with Pi, DHAP, or 3PGA (Borchert et al., 1989, 1993). The Vmax value for Glc6P uptake, using the release of 32P-labelled Pi to measure back exchange, was measured at 0.51 µmol mg–1 protein h–1. Transport of Glc6P across the plastid membrane could limit flux of hexose phosphate through the oxidative pentose phosphate pathway. DIDS was found to be a potent inhibitor of Glc6P-dependent nitrite reduction (Fig. 1). The nature of the covalent modification means it was not possible to measure the direct effects of DIDS on Glc6P uptake, therefore this experiment was repeated using known physiological competitive inhibitors of Glc6P uptake, Pi, PGA or DHAP (Fig. 2). It was found that even when approximately 50% of Glc6P uptake was inhibited by Pi, PGA, or DHAP, sufficient Glc6P was imported to support nitrite reduction (Fig. 3). Although these experiments are in vitro studies, it seems likely that control over Glc6P-dependent nitrite reduction is not through the transport of Glc6P. Since the GPT catalyses the counter exchange of Glc6P and DHAP, it is perhaps not surprising that the GPT exerts such little control over Glc6P-dependent nitrite reduction. A high control coefficient for Glc6P transport would mean that the process of nitrite reduction within the organelle could otherwise be severely limited by the presence of Pi, triosephosphate, and PGA in the cytosol.
The uptake of inorganic nitrite into the plastids is another possible point of regulation of Glc6P-dependent nitrite reduction. Nitrite moves from the cytosol to the plastid where it is reduced by NiR to ammonium. Since nitrite is toxic, the intracellular concentration is kept at a low level in normal circumstances through a combination of NR expression, NR catalytic activity, and NR protein degradation (Kaiser and Huber, 2001). When the nitrite reducing capacity is low, protein phosphorylation inactivates NR (Kaiser and Huber, 2001). Nitrite may be transported freely in its protonated form as nitrous acid or as an ion. In plants the molecular mechanism of nitrite uptake into plastids is unclear and there is conflicting data regarding the presence of a specific nitrite transporter (Brunswick and Cresswell, 1988; Shingles et al., 1996). In Chlamydomonas reinhardtii, the unicellular eukaryotic and photosynthetic green alga, plastidic nitrite transporters allowing the entry of nitrite into the chloroplasts have been identified (Mariscal et al., 2004; Rexach et al., 2000). To date, no potential nitrite transporters have been identified in higher plants. In the experiments reported here the availability of 13N-labelled nitrite has meant that it has been possible to perform plastid labelling studies and determine uptake of nitrite into plastids over a range of nitrite concentrations. It is clear from Fig. 4 that nitrite enters the root plastids by diffusion and as such is unlikely to regulate Glc6P-dependent nitrite reduction. Indeed, its transport across the plastid envelope increases if nitrite is being reduced by NiR (data not shown).
It has previously been shown that within pea root plastids, the oxidation of Glc6P by the OPPP is capable of supporting the activity of NiR (Bowsher et al., 1989) and GOGAT (Bowsher et al., 1992). Furthermore, from the studies reported here, under substrate saturating conditions, NiR and GOGAT compete for reductant-generated by the OPPP (Fig. 5). The restriction on Glc6P-dependent nitrogen metabolism appears to be due to the rate of supply of reductant as opposed to the ability of enzymes of nitrogen assimilation to utilize it. The reductant supply may be restricted in a number of possible ways. Of all the OPPP enzymes, it is G6PDH which is usually found to have the lowest maximal activity (Emes and Fowler, 1983). Depending on status of nitrate induction, pea root G6PDH activity was 2556–8150 nmol mg–1 protein h–1. To support the maximum rate of nitrite reduction of 1000 nmol mg–1 protein h–1 (Fig. 5b), 1500 nmol mg–1 protein h–1 Glc6P would need to be oxidized. To support the maximum rate of glutamate formation of 1500 nmol mg–1 protein h–1 (Fig. 5c), 375 nmol mg–1 protein h–1 of Glc6P would need to be oxidized. The total amount of Glc6P which would need to be oxidized to support both nitrite reduction and glutamate synthesis under saturating conditions is thus 1875 nmol mg–1 protein h–1, indicating that the capacity of G6PDH is more than sufficient to meet this demand and nearly 6-fold in excess when fully induced. G6PDH has been reported to be strongly inhibited by a high NADPH/NADP ratio (Wright et al., 1997). Studies on isolated barley root plastids suggest that it is the P2 isoform of G6PDH which is present in root plastids (Esposito et al., 2001a). This isoform appears able to continue to support reactions under NADPH/NADP ratios that would strongly inhibit the chloroplastic and cytosolic isoforms of the enzyme (Wright et al., 1997; Esposito et al., 2001a) indicating that the ratio of reduced to oxidized pyridine nucleotide is also unlikely to be a limitation.
The oxidation of 1 mol of Glc6P via G6PDH and 6PGDH generates four electrons (2 NAPDH). To achieve electron transfer efficiently from the OPPP to the enzymes of nitrite assimilation under saturating conditions, 1870 nmol mg–1 protein h–1 would require FNR to operate at a rate of 7632 nmol mg–1 protein h–1. Previously, it has been reported that an FNR activity of 9000–18 000 nmol mg–1 protein h–1 was in root plastids isolated from plants induced with 10 mM nitrate (Bowsher et al., 1993). This rate was determined using a non-physiological cytochrome c-based assay and represents a one electron transfer. Although the maximum rate of transfer of NADPH to ferredoxin is not known, if it is assumed to be equivalent to the rate of cytochrome c reduction, FNR may not limit flux through the OPPP. However, since this is only slightly in excess of the rate required to support nitrite reduction and glutamate synthesis, it is an area worthy of further investigation. An alternative point of control may be competition between NiR and GOGAT for reduced ferredoxin. Yonekura-Sakibara et al. (2000) found that maize roots contained 0.41 µg (approximately 410 nmol) mg–1 protein ferredoxin, and 1.08 µg (approximately 180 nmol) mg–1 protein sulphite reductase (another ferredoxin-dependent bioassimilatory enzyme). Such equivalent concentrations of the electron donor and only one of its dependent enzymes raises the possibility that access to unbound, reduced ferredoxin may limit the activity of several enzymes of reductive assimilation, including NiR and GOGAT.
The results described here suggest that the restriction on Glc6P-dependent nitrogen metabolism in root plastids is due to the rate of supply of reductant as opposed to the ability of the enzymes of nitrogen assimilation to accept reductant. Competition for electrons between nitrite reductase and glutamate synthase imply that the OPPP is not capable of producing sufficient reductant to support the optimal activities of these two enzymes simultaneously. Although NiR and GOGAT represent different sink strengths needing six electrons and two electrons per reaction, respectively, they inhibit each other approximately in proportion and the percentage reduction in activity during competition for reductant is approximately the same for each enzyme (Fig. 7). For example, at a concentration of 1 mM Glc6P, nitrite reduction is decreased by approximately 20% in the presence of saturating glutamine and 2-oxoglutarate. Likewise, at the same Glc6P concentration, GOGAT activity is decreased by 20% in the presence of saturating nitrite. This suggests that the oxidation of Glc6P through the OPPP may place a significant constraint on the reactions of nitrogen assimilation in plastids. In addition, there may be a mechanism for partitioning of electrons between the two enzymes so that they remain in balance. Such partitioning may be essential to the functioning of nitrogen assimilation in vivo. An additional consideration must be given to species where the root is the principal site of nitrate reduction. In Lotus japonicus, for example, as with many other temperate legumes, there must be a heavy demand for electrons from carbohydrate oxidation which extends throughout the diurnal cycle (Prosser et al., 2006).
|
Nitrate assimilation is a highly regulated and integrated metabolic process (Stitt, 1999). It is dependent on a range of signals including the amount and type of nitrogen source, light and carbon availability. The complex nature of this regulation reflects the multiple levels of control. Non-photosynthetic plastids are involved in a range of other metabolic processes including fatty acid synthesis. If other pathways are superimposed onto the reductant demands of nitrogen assimilation, carbohydrate oxidation within plastids is likely to become even more limiting for any given activity and thus play an important role in regulating assimilatory and biosynthetic pathways.
|
Acknowledgements |
|---|
The authors are grateful to the BBSRC, MWO, and Royal Society for financial support. They also gratefully acknowledge the PET Center, Groningen, for providing 13N.
|
Abbreviations |
|---|
DHAP, dihydroxyacetone phosphate; DIDS, 4, 4'-diisothiocyanatostilbene-2-2'-disulphonic acid; G6PDH, glucose-6-phosphate dehydrogenase; Glc6P, glucose-6-phosphate; Gln, glutamine; Glu, glutamate; GOGAT, glutamate synthase; GS, glutamine synthetase; GPT, Glc6P/phosphate translocator; HP, hexose phosphates; NiR, nitrite reductase; OPPP, oxidative pentose phosphate pathway; PEP, phosphoenolpyruvate; PPT, phosphoenolpyruvate/phosphate translocator; Pi, inorganic phosphate; PGA, phosphoglyceric acid; TPT, triose phosphate/phosphate translocator.
|
References |
|---|
|
TopAbstractIntroductionMaterials and methodsResultsDiscussionReferences |
|---|
Andrews M. (1986) The partitioning of nitrate assimilation between root and shoot of higher plants. Plant, Cell and Environment 9 511–519.
Andrews M, Lea PJ, Raven JA, Lindsey K. (2004) Can genetic manipulation of plant nitrogen assimilation enzymes result in increased crop yield and greater N-use efficiency? An assessment. Annals of Applied Biology 145 25–40.[CrossRef][Web of Science]
Borchert S, Große H, Heldt HW. (1989) Specific transport of inorganic phosphate, glucose 6-phosphate, dihydroxyacetone phosphate, and 3-phosphoglycerate into amyloplasts. FEBS Letters 253 183–186.[CrossRef][Web of Science]
Borchert S, Harborth J, Schünemann D, Hoferickter WP, Heldt HW. (1993) Studies of the enzymic capacities and transport properties of pea root plastids. Plant Physiology 101 303–312.[Abstract]
Bowsher CG, Hucklesby DP, Emes MJ. (1993) Induction of ferredoxin-NADP+ oxidoreductase and ferredoxin synthesis in pea root plastids during nitrate assimilation. The Plant Journal 3 463–467.[Web of Science]
Bowsher CG, Boulton EL, Rose J, Nayagam S, Emes MJ. (1992) Reductant for glutatmate synthase is generated by the oxidative pentose phosphate pathway in non-photosynthetic root plastids. The Plant Journal 2 893–898.[CrossRef][Web of Science]
Bowsher CG, Hucklesby DP, Emes MJ. (1989) Nitrite reduction and carbohydrate metabolism in plastids from roots of Pisum sativum L. Planta 177 359–366.[CrossRef][Web of Science]
Bradford MM. (1976) A rapid and sensitive method for quantitation of microgram quantities of protein utilizing the principle of protein-dye binding. Analytical Biochemistry 72 248–254.[CrossRef][Web of Science][Medline]
Brewer PG, Chan K, Riley JP. (1965) Automatic determination of certain micronutrients in sea water. In Skeggs LT (Ed.). Automation of analytical chemistry. Technicon Symposium pp. 303–314.
Brunswick P and Cresswell CF. (1988) Nitrite uptake into intact pea chloroplasts. I. Kinetics and relationship with nitrite assimilation. Plant Physiology 86 378–383.
Crawford NM. (1995) Nitrate: nutrient and signal for plant growth. The Plant Cell 7 859–868.[CrossRef][Web of Science][Medline]
Debnam PM and Emes MJ. (1999) Subcellular distribution of enzymes of the oxidative pentose phosphate pathway in root and leaf tissues. Journal of Experimental Botany 50 1653–1651.
Emes MJ and England S. (1986) Purification of plastids from higher-plant roots. Planta 168 161–166.[Web of Science]
Emes MJ and Fowler MW. (1979) The intracellular location of the enzymes of the nitrate assimilation in the apices of seedling pea roots. Planta 144 249–253.[CrossRef][Web of Science]
Emes MJ and Fowler MW. (1983) The supply of reducing power for nitrate reduction in plastids of seedlings of pea roots (Pisum sativum L.). Planta 158 97–102.[CrossRef][Web of Science]
Emes MJ and Traska A. (1987) Uptake of inorganic phosphate by plastids purified from the roots of Pisum sativum L. Journal of Experimental Botany 38 1781–1788.
Esposito S, Carfagna S, Massaro G, Vona V, Di Martino Rigano V. (2001a) Glucose-6P dehydrogenase in barley roots: kinetic properties and localization of the isoforms. Planta 212 627–634.[CrossRef][Web of Science][Medline]
Esposito S, Massaro G, Vona V, Di Martino Rigano V, Carfagna S. (2001b) Ammonium induction of a novel isoform of glucose-6P dehydrogenase in barley roots. Physiologia Plantarum 113 469–476.[CrossRef]
Esposito S, Massaro G, Vona V, Di Martino Rigano V, Carfagna S. (2003) Glutamate synthesis in barley roots: the role of the plastidic glucose-6-phosphate dehydrogenase. Planta 216 639–647.[CrossRef][Web of Science][Medline]
Fischer K, Kammerer B, Gutensohn M, Arbinger B, Weber RE, Hausler RE, Flugge UI. (1997) A new class of plastidic phosphate translocators: a putative link between primary and secondary metabolism by the phosphoenolpyruvate/phosphate antiporter. The Plant Cell 9 453–462.[Abstract]
Fliege R, Flügge UI, Werden K, Heldt HW. (1978) Specific transport of inorganic phosphate, 3-phosphoglycerate and triosephosphates across the inner membrane of the envelope in spinach chloroplasts. Biocimica et Biophysica Acta 502 232–247.[CrossRef]
Flugge UI. (1999) Phosphate translocators in plastids. Annual Review of Plant Physiology and Plant Molecular Biology 50 27–45.[CrossRef][Web of Science][Medline]
Fowler MW and ap Rees T. (1970) Carbohydrate oxidation during differentiation in roots of Pisium sativum. Biochimica et Biophysica Acta 201 33–44.[Medline]
Good AG, Shrawat AK, Muench DG. (2004) Can less yield more? Is reducing nutrient input into the environment compatible with maintaining crop production? Trends in Plant Science 9 597–605.[CrossRef][Web of Science][Medline]
Grasshoff K. (1964) Zur Bestimmung von Nitrat in Meer- und Trinkwasser Keiler Meeresforsch. Bs 20(No 1) 5–11.
Hartwell J, Bowsher CG, Emes MJ. (1996) Purification of plastids from higher-plant roots. Planta 168 161–166.
Heldt HW. (1980) Measurement of metabolite movement across the envelope and of the pH in the stroma and the thylakoid space in intact chloroplasts. Methods in Enzymology 69 604–613.
Kaiser WM and Huber SC. (2001) Post-translational regulation of nitrate reductase: mechanism, physiological relevance and environmental triggers. Journal of Experimental Botany 52 1981–1989.
Kammerer B, Fischer K, Hilpert B, Schubert S, Gutensohn M, Weber A, Flugge U-I. (1998) Molecular characterization of a carbon transporter in plastids from heterotrophic tissues: the glucose 6-phosphate/phosphate antiporter. The Plant Cell 10 105–117.
Knappe S, Flugge U-I, Fischer K. (2003) Analysis of the plastidic phosphate translocator gene family in Arabidopsis and identification of new phosphate translocator-homologous transporters, classified by their putative substrate-binding site. Plant Physiology 131 1178–1190.
Knight JS and Gray JC. (1994) Expression of genes encoding the tobacco chloroplast phosphate translocator is not light-regulated and is repressed by sucrose. Molecular and General Genetics 242 586–594.[CrossRef]
Knight JS, Emes MJ, Debnam PM. (2001) Isolation and characterization of a full-length genomic clone encoding a plastidic glucose 6-phosphate dehydrogenase from Nicotiana tabacum. Planta 212 499–507.[CrossRef][Web of Science][Medline]
Kruger NJ and Von Schaewen A. (2003) The oxidative pentose phosphate pathway: structure and organization. Current Opinion in Plant Biology 6 236–246.[CrossRef][Web of Science][Medline]
Lam H-M, Coschigano KT, Oliveira IC, Melo-Oliveira R, Coruzzi GM. (1996) The molecular-genetics of nitrogen assimilation into amino acids in higher plants. Annual Review of Plant Physiology and Plant Molecular Biology 47 569–593.[CrossRef][Web of Science][Medline]
Mariscal V, Rexach J, Fernandez E, Galván A. (2004) The plastidic nitrite transporter NARI;I improves nitrate use efficiency for growth in Chlamydomanas. Plant, Cell and Environment 27 1321–1328.[CrossRef]
Matsumara T, Sakakibara H, Nakano R, Kimata Y, Sugiyama T, Hase T. (1997) A nitrate-inducible ferredoxin in maize roots: genomic organization and differential expression of two non-photosynthetic ferredoxin isoproteins. Plant Physiology 114 653–660.[Abstract]
Niewiadomski P, Knappe S, Geimer S, Fischer K, Schulz B, Unte US, Rosso MG, Ache P, Flugge U-I, Schneider A. (2005) The Arabidopsis plastidic glucose 6-phosphate/phosphate translocator GPT1 is essential for pollen maturation and embryo sac development. The Plant Cell 17 760–775.
Pate JS. (1980) Transport and partitioning of nitrogenous solutes. Annual Review of Plant Physiology 31 313–340.[Web of Science]
Prosser IM, Massonneau A, Smyth AJ, Waterhouse RN, Forde BG, Clarkson DT. (2006) Nitrate assimilation in the forage legume Lotus japonicus L. Planta 223 821–834.[CrossRef][Web of Science][Medline]
Redinbaugh MG and Campbell WH. (1998) Nitrate regulation of the oxidative pentose phosphate pathway in maize (Zea mays L.) root plastids: induction of 6-phosphogluconate dehydrogenase activity, protein and transcript levels. Plant Science 134 129–140.
Rexach J, Fernandez E, Galvan A. (2000) The Chlamydomonas reinhardtii Nar1 gene encodes a chloroplast membrane protein involved in nitrite transport. The Plant Cell 12 1441–1453.
Ritchie SW, Redinbaugh MG, Shiraishi N, Vrba Kohler JM, Campbell WH. (1994) Identification of a maize root transcript expressed in the primary response to nitrate: characterization of a cDNA with homology to ferredoxin-NADP+ oxidoreductase. Plant Molecular Biology 26 679–690.[CrossRef][Web of Science][Medline]
Schnarrenberger C, Flechner A, Martin W. (1995) Enzymatic evidence for a complete oxidative pentose phosphate pathway in chloroplasts and on in complete pathway in the cytosol of spinach leaves. Plant Physiology 108 609–614.[Abstract]
Shingles R, Roh MH, McCarty RE. (1996) Nitrite transport in chloroplast inner envelope vesicles. 1. Direct measurement of proton-linked transport. Plant Physiology 112 1375–1381.[Abstract]
Stitt M. (1999) Nitrate regulation of metabolism and growth. Current Opinion in Plant Biology 2 178–196.[CrossRef][Web of Science][Medline]
Wakao S and Benning C. (2005) Genome-wide analysis of glucose-6-phophate dehydrogenase in Arabidopsis. The Plant Journal 41 243–256.[CrossRef][Web of Science][Medline]
Wang RC, Guegler K, LaBrie ST, Crawford NM. (2000) Genomic analysis of a nutrient response in arabidopsis reveals diverse expression patterns and novel metabolic and potential regulatory genes induced by nitrate. The Plant Cell 12 1491–1509.
Wang Y-H, Garvin DF, Kochian LV. (2001) Nitrate-induced genes in tomato roots. Array analysis reveals novel genes that may play a role in nitrogen nutrition. Plant Physiology 127 345–359.
Wood ED, Armstrong FAJ, Richards FA. (1967) Determination of nitrate in sea water by cadmium-copper reduction to nitrate. Journal of Marine Biology 47 23–31.
Wright DP, Huppe HC, Turpin DH. (1997) In vivo and in vitro studies of glucose-6-phosphate dehydrogenase from barley root plastids in relation to reductant supply for assimilation. Plant Physiology 114 1413–1419.[Abstract]
Yonekura-Sakakibara K, Onda Y, Ashikari T, Tanaka Y, Kusumi T, Hase T. (2000) Analysis of reductant supply systems for ferredoxin-dependent sulfite reductase in photosynthetic and non-photosynthetic organs of maize. Plant Physiology 122 887–894.
CiteULike 
Connotea 
Del.icio.us What's this?
This article has been cited by other articles:
|
|
 |
|
  Y.-J. Liu, A. Nunes-Nesi, S. V. Wallstrom, I. Lager, A. M. Michalecka, F. E.B. Norberg, S. Widell, K. M. Fredlund, A. R. Fernie, and A. G. Rasmusson A Redox-Mediated Modulation of Stem Bolting in Transgenic Nicotiana sylvestris Differentially Expressing the External Mitochondrial NADPH Dehydrogenase Plant Physiology, July 1, 2009; 150(3): 1248 - 1259. [Abstract] [Full Text] [PDF]
|
|
|
|
 |
|
 B. G. Forde and P. J. Lea Glutamate in plants: metabolism, regulation, and signalling J. Exp. Bot., July 1, 2007; 58(9): 2339 - 2358. [Abstract] [Full Text] [PDF]
|
|
| ||||||||||||||||||||||||||||||||||||||||||||||||||||||||||||||||||||||||||||||||||||