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Keywords:

  • day respiration;
  • glutamate;
  • isotopes;
  • labelling;
  • nitrogen assimilation

Summary

  1. Top of page
  2. Summary
  3. Introduction
  4. Materials and Methods
  5. Results
  6. Discussion
  7. Acknowledgements
  8. References
  9. Supporting Information
  • Nitrogen assimilation in leaves requires primary NH2 acceptors that, in turn, originate from primary carbon metabolism. Respiratory metabolism is believed to provide such acceptors (such as 2-oxoglutarate), so that day respiration is commonly seen as a cornerstone for nitrogen assimilation into glutamate in illuminated leaves. However, both glycolysis and day respiratory CO2 evolution are known to be inhibited by light, thereby compromising the input of recent photosynthetic carbon for glutamate production.
  • In this study, we carried out isotopic labelling experiments with 13CO2 and 15N-ammonium nitrate on detached leaves of rapeseed (Brassica napus), and performed 13C- and 15N-nuclear magnetic resonance analyses.
  • Our results indicated that the production of 13C-glutamate and 13C-glutamine under a 13CO2 atmosphere was very weak, whereas 13C-glutamate and 13C-glutamine appeared in both the subsequent dark period and the next light period under a 12CO2 atmosphere. Consistently, the analysis of heteronuclear (13C–15N) interactions within molecules indicated that most 15N-glutamate and 15N-glutamine molecules were not 13C labelled after 13C/15N double labelling. That is, recent carbon atoms (i.e. 13C) were hardly incorporated into glutamate, but new glutamate molecules were synthesized, as evidenced by 15N incorporation.
  • We conclude that the remobilization of night-stored molecules plays a significant role in providing 2-oxoglutarate for glutamate synthesis in illuminated rapeseed leaves, and therefore the natural day : night cycle seems critical for nitrogen assimilation.

Introduction

  1. Top of page
  2. Summary
  3. Introduction
  4. Materials and Methods
  5. Results
  6. Discussion
  7. Acknowledgements
  8. References
  9. Supporting Information

It is now well recognized that plant growth and development depend on the interaction between carbon and nitrogen metabolic pathways. Indeed, optimal rates of plant biomass production and photosynthesis (CO2 fixation) require adequate nitrogen supply, which is, in turn, associated with CO2 losses through respiration for nitrate absorption, reduction and assimilation to organic compounds (for a review, see Lawlor, 2002). Carbon–nitrogen interactions are therefore a cornerstone for plant productivity, and a better understanding of the metabolic basis of such interactions is critical for future improvements (Lawlor, 2002).

In illuminated leaves, nitrogen reduction and assimilation involve the operation of nitrate and nitrite reductase (EC 1.6.6.1 and EC 1.7.7.1) and the glutamine synthetase/glutamine-2-oxoglutarate aminotransferase (GS/GOGAT, EC 6.3.1.2 EC 1.4.1.13) pathway, which produces glutamate (Glu) for other amino acid biosynthesis (for a recent review, see Forde & Lea, 2007). The regulation of GS and nitrate reductase activity (e.g. the availability of reductants) is such that leaf nitrogen assimilation is mainly achieved in the light relative to the dark as a result of the requirement for ATP and reductants of several nitrogen-assimilatory enzymes (Delhon et al., 1995; Stitt et al., 2002). Roots are nevertheless responsible for a variable, species-specific proportion of nitrate reduction either in the dark or in the light (Radin, 1977, 1978). 15N-Isotopic labelling has further shown that nitrate molecules that are not consumed by roots in the dark are exported to leaves (shoots), where they accumulate and become available for reduction during the subsequent light period (Gojon et al., 1986). Although leaf nitrate content is ordinarily large (Gojon et al., 1986), thereby allowing isotopic dilution and somewhat impeding 15N labelling, some nitrogen recycling (e.g. protein hydrolysis) is thought to occur in leaf cells (Bauer et al., 1977; and see Tcherkez & Hodges, 2008 for a review).

By contrast, the origin of carbon atoms (in the form of 2-oxoglutarate) for Glu and glutamine (Gln) production is currently much less well characterized. Day respiration may play a key role in supplying carbon skeletons through the operation of the tricarboxylic acid (TCA) cycle and, accordingly, it has been shown that the day respiration rate Rd (respiratory CO2 evolution in the light) is sensitive to nitrogen assimilation (Guo et al., 2005). In addition, calculations based on leaf citrate content available at the beginning of the light period suggest that it is not sufficient to feed 2-oxoglutarate synthesis for Glu production (Stitt et al., 2002). Therefore, day respiration might be critical for supplementing 2-oxoglutarate synthesis, possibly accompanied by the anaplerotic activity of phosphoenolpyruvate carboxylase (PEPc, EC 4.1.1.31; Huppe & Turpin, 1994). However, it is well accepted that leaf respiration is inhibited by light (for a review, see Atkin et al., 2000) and is associated with a restricted flux through TCA cycle enzymes (Hanning & Heldt, 1993; Tcherkez et al., 2005). In addition, mutants affected in either aconitase (EC 4.2.1.3) or isocitrate dehydrogenase (EC 1.1.1.41 and EC 1.1.1.42) activity do not show any significant reduction in plant biomass or nitrogen content (Kruse et al., 1998; Carrari et al., 2003; Lemaitre et al., 2007). Such conflicting results indicate that presumably the carbon source that feeds Glu production may consist of both newly synthesized (TCA-derived) and remobilized (e.g. derived from night citrate) 2-oxoglutarate molecules. Accordingly, 14C-labelling and radiometric studies of day-evolved CO2 have suggested that up to 40% of decarboxylated CO2 comes from stored, low turned-over carbon molecules (Parnik et al., 2002). Nevertheless, to our knowledge, no metabolic study has both demonstrated and quantified the contribution of current CO2 assimilation and storage remobilization as carbon sources for Glu synthesis.

The aim of this study was to determine the contribution of current fixed and stored carbon to Glu synthesis. To address this objective, we investigated the synthesis of major metabolites (Glu and respiratory intermediates) in the light using double 13C and 15N labelling. Double isotopic labelling has already been used at the plant level to estimate nitrogen partitioning from the natural isotopic composition of total organic matter (see, for example, Maillard et al., 1994), but, to our knowledge, the metabolic pathways after double labelling have never been examined. In this study, illuminated leaves of rapeseed (Brassica napus) were labelled with 13CO2 and 15N-ammonium nitrate, and subsequently analyzed with 13C- and 15N-nuclear magnetic resonance (NMR), to trace 13C and 15N atoms within metabolites and to detect double-labelled molecules (i.e. simultaneous 13C and 15N within single molecules) characterized by spin–spin interactions. Our results showed that the majority of 15N-Glu and 15N-Gln molecules were not 13C labelled, whereas up to one-half of 13C-Glu or 13C-Gln pools were also 15N labelled. This discrepancy indicates that a significant part of the carbon skeleton that is derived from current assimilation through day respiration is used for nitrogen assimilation, whereas such a carbon input is quantitatively modest with respect to total nitrogen assimilation.

Materials and Methods

  1. Top of page
  2. Summary
  3. Introduction
  4. Materials and Methods
  5. Results
  6. Discussion
  7. Acknowledgements
  8. References
  9. Supporting Information

Plant material

Seeds of rape plants (Brassica napus L. var. oleifera cv. Darmor) were germinated in Petri dishes on filter paper. After 72 h, seedlings were transferred to 500 ml pots filled with potting mix. Plants were grown in the glasshouse under 22°C : 18°C, 60% : 55% relative humidity, 16 h : 8 h photoperiod (day : night) as described by Vartanian et al. (1987). Plants were automatically watered three times a day with nutritive solution (Hydrokani C2, Yara, Chambourcy, France) in which the ammonium nitrate was checked to be at natural 15N abundance (δ15N = +2.69 ± 0.61‰, where δ15N is the nitrogen isotope composition with respect to atmospheric N2), that is, 0.37%. Carbon dioxide in air was at natural 13C abundance (δ13C = −8.92 ± 0.55‰, where δ13C is the carbon isotope composition with respect to V-PDB), that is, 1.1%.

Chemicals

Labelling experiments were carried out with 13CO2 99% (Air Liquide, Grigny, France). Ordinary CO2 (at near-natural abundance, that is, 1.1%) had a δ13C value of −50.05 ± 0.49‰ (Air Liquide). Feeding solutions supplied to detached leaves were all at 15 mmol l−1 ammonium nitrate, either 99% enriched in 15N (Isotec-Sigma-Aldrich, Saint Quentin Fallavier, France) or at natural abundance (δ15N = −0.18 ± 0.06‰) (Prolabo, Fontenay-sous-Bois France). In all cases, the pH was adjusted to pH 7 with HCl.

Labelling methods and gas exchange

In all experiments, fifth-ranked leaves (from the apex) on 6-wk-old plants were used (average surface area of 90 cm2). After 7 h light in the glasshouse, a leaf was cut under water and transferred to darkness for 30 min. It was then placed in a purpose-built assimilation chamber connected to the sample tube of an Li-6400 open system (LiCor, Lincoln, Nebraska, USA), as already described by Tcherkez et al. (2005). Each leaf was illuminated for 1 h [400 μmol m−2 s−1 photosynthetically active radiation (PAR), 21°C, 21% O2] with CO2 at natural 13C abundance (1.1%13C) to reach the photosynthetic steady state. Each leaf was then simultaneously fed with 15N-ammonium nitrate at the same time that CO2 was changed to 99%13CO2. The duration of labelling was 1, 6 or 12 h, after which the leaf was immediately frozen in liquid nitrogen. In another set of experiments, 6 h light in 13CO2 was followed by either 4 h darkness, or 4 h darkness followed by 6 h in CO2 at natural 13C abundance.

During the experiments, the CO2 mole fraction in air (ca) was maintained at either 400 μmol mol−1 (ordinary conditions) or 100 μmol mol−1 (highly photorespiratory conditions). Identical experiments were carried out using only CO2 and ammonium nitrate at natural isotopic abundances, as NMR controls (for 13C- and 15N-enrichment calculations, see below). All samples were kept at −80°C before extraction for NMR analyses.

NMR

Perchloric acid extracts were prepared from 2.5 g of frozen leaf material, as described by Aubert et al. (1996) for phloem cells and applied in Tcherkez et al. (2005) for leaves. Both 15N- and 13C-NMR spectra were recorded using a Bruker NMR spectrometer (AMX 400, wide bore, Bruker, Wissenbourg, France) equipped with a 10 mm multinuclear probe tuned at 40.55 MHz.

13C-NMR acquisition was carried out with 19 μs pulses (90°) at 6 s intervals and a sweep width of 20 kHz. Broad-band decoupling at 2.5 W during acquisition and 0.5 W during the delay was applied using Waltz sequences; the signal was digitized using 32 000 data points zero-filled to 64 000 and processed with 0 Hz line broadening. 13C-NMR spectra were referenced to hexamethyldisiloxane at 2.7 ppm Mn2+ ions were chelated by the addition of 1 mmol l−1 1,2-cyclohexylenedinitrilotetraacetic acid. The assignment of resonance of 13C peaks was carried out according to Gout et al. (1993).

15N-NMR acquisition was carried out with 8 μs pulses (70°) at 2 s intervals and a spectral width of 16 kHz. Broad-band decoupling at 10 W during acquisition and 0.5 W during delay was applied using Waltz sequences; the signal was digitized using 64 000 data points zero-filled to 128 000 and processed with either 0.5 or 0.2 Hz exponential line broadening. 15N-NMR spectra were referenced to 15N-ammonium chloride (pH 7.5 in H2O/D2O 20%) at 0 ppm. The identified compounds were quantified from the area of the resonance peaks.

In the case of spin–spin interaction between 15N and 13C atoms or between 13C atoms, isotopic signals were considered as the sum of multiplets. In both 13C and 15N techniques, peak intensities were normalized to a known amount of an internal reference compound (500 μmol maleate for 13C-NMR and 5 μmol β-alanine for 15N-NMR), which was added to the sample during extraction. Isotopic enrichments were calculated using the relative isotopic signal (13C or 15N signals normalized by the reference compound) obtained after labelling divided by that obtained after a control experiment (natural abundance) and then multiplied by the natural abundance (1.1% for 13C and 0.37% for 15N). For 15N-enrichment calculations, any change in the transpiration rate (leaf absorption of ammonium nitrate through the transpiration stream) was taken into account by multiplying by the transpiration ratio between the 15N experiment and that carried out at natural abundance. 13C–15N spin–spin interactions were quantified (in %) by dividing the signal corresponding to interaction peak(s) by the total isotopic signal.

Clustering analysis

The 13C-NMR data were represented as an isotopomic array, as described in Tcherkez et al. (2007). The positional isotopic abundances (in 13C %) relative to the natural 13C abundance (1.1%) are indicated by colours, so that black cells indicate near-natural abundance, and green and red cells indicate lower and higher than natural 13C abundance, respectively. The clustering analysis was carried out with MeV 4.1 software (Saeed et al., 2003) and based on the cosine correlation method.

Results

  1. Top of page
  2. Summary
  3. Introduction
  4. Materials and Methods
  5. Results
  6. Discussion
  7. Acknowledgements
  8. References
  9. Supporting Information

13C assimilation and tracing in metabolites

CO2 fixation was monitored with a gas exchange system, and photosynthetic assimilation, transpiration and internal CO2 concentration were measured concurrently (the values are shown in Fig. 1). There was a decrease of approximately four-fold in the CO2 assimilation rate under highly photorespiratory conditions (Fig. 1a). This agreed with the observed decrease in the internal CO2 mole fraction (300 to 75 μmol mol−1) between the normal and high photorespiratory conditions, respectively (Fig. 1c). Stomatal conductance was observed to be slightly higher at 100 μmol mol−1 CO2, and therefore the absorption of ammonium nitrate through the transpiration stream was also higher. This difference was taken into account when calculating the relative 15N content of Glu (see below, Fig. 2).

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Figure 1.  Net photosynthetic assimilation (a), H2O transpiration (b) and intercellular CO2 concentration (ci, c) of detached Brassica napus leaves during isotopic labelling with [13C]-CO2 and [15N]-NH4NO3 under either 100 μmol mol−1 CO2 (filled bars) or 400 μmol mol−1 CO2 (open bars), 21% O2, 400 μmol m−2 s−1 photosynthetically active radiation (PAR) and 21°C.

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image

Figure 2.  Relationship between the relative 15N commitment and 13C commitment of absorbed 15N and 13CO2 into glutamate (Glu) in Brassica napus leaves, calculated from the 13C- and 15N-NMR enrichments in Glu (Fig. 3) and gas exchange (Fig. 1) data. Each datum corresponds to the mean value for a given labelling time; that is, the labelling time increases from left to right, as indicated by the superscripts 1, 6 and 12 (corresponding to hours) under 100 μmol mol−1 CO2 (closed circle) or 400 μmol mol−1 CO2 (open circle), 21% O2, 400 μmol m−2 s−1 PAR and 21 °C.

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The 13C enrichment of metabolites is shown in Fig. 3, in which red cells represent 13C abundances above natural abundance. Four different major groups were obtained with the hierarchical clustering analysis (Fig. 3, left). Regardless of such groups, the most labelled metabolites (major 13C sinks under our experimental conditions) were glycine (Gly), serine (Ser), malate (Mal), aspartate (Asp), alanine (Ala), valine, Glu and Gln. Such a pattern was not very sensitive to photorespiration, except for the values of the first group.

image

Figure 3.  Isotopomics array representation of the 13C enrichment in C-atom positions detected by 13C-NMR after isotopic labelling of detached Brassica napus leaves. Labelling conditions were (raws in that order): 1 h, 6 h, 12 h of 13CO2 labelling, 6 h of 13CO2 labelling followed by 30 min in darkness (indicated as 6 h + 30), 6 h of 13CO2 labelling followed by 4 h in darkness and then 6 h with natural CO2 (1.1%13CO2) (indicated as 6/4/6 h). The CO2 mole fraction was either 100 or 400 μmol mol−1 (indicated as 100 and 400 on raw titles) and the gas exchange conditions were 21% O2, 400 μmol m−2 s−1 photosynthetically active radiation (PAR) and 21°C. Ala, alanine; Asp, aspartate; Cit, citrate; Ctr-Me, one methyl of citrulline; GABA, γ-aminobutyrate; Glc, glucose; Gln, glutamine; Glu, glutamate; Gly, glycine; Gth, glutathione; Homo-Ser, homo-serine; Icit, isocitrate; Ile, isoleucine; Leu, leucine; Lys, lysine; Mal, malate; Met, methionine; Obut, oxobutyrate; PGA, phosphoglycerate; Pyr, pyruvate; Ser, serine; Succ, succinate; Thr, threonine; Val, valine; Val-Me1, one of the two methyl groups of valine; Val-Me2, the other methyl group of valine. Carbon atom positions are indicated by numbers (international chemical nomenclature). In each cell, the colour stands for the positional 13C abundance (green, < 1.1%13C; red, > 1.1%13C). Left hand side: clustering tree obtained with the cosine correlation of 13C enrichments.

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The C-3 and C-4 atom positions in Glu and C-1 and C-5 positions in Gln were all within the first group (I), which also included the C-2 and C-3 atom positions of Asp and Mal, thereby suggesting that the PEPc-catalysed carboxylation of 13C-enriched phosphoenolpyruvate occurred, and this, in turn, labelled molecules downstream, such as Glu, via the TCA cycle (see also the metabolic scheme in Fig. S3, Supporting Information). Accordingly, the 13C-labelling pattern in the C-2 atom position in Glu and Gln covaried with that in C-1 or C-4 in Mal (group IV). In addition, the very similar labelling in C-1 and C-4 in Mal indicated that the Mal/fumarate backward equilibrium took place and redistributed the PEPc-derived 13C label in C-4 into the C-1 position. Citrate C-3 appeared labelled and clustered with the latter Mal positions, whereas the C-1 and C-5 atom positions were hardly labelled. This observation can be accounted for by the isotopic dilution of 13C enrichment through glycolysis and the diversion of the C-2 atom position of pyruvate into Ala, which was indeed strongly 13C enriched. As a result, most of the 13C label found in citrate was that originating from phosphoenolpyruvate through PEPc and citrate synthase. It is noteworthy that succinate was always poorly labelled and only exhibited weak labelling in C-2/C-3 when the 13CO2 labelling period in the light was followed by 4 h darkness and 6 h in the light with 12CO2 (right-most raw). This suggests that succinate molecules were not labelled by 13CO2 in the light under our conditions and that the synthesis of 13C-succinate required the onset of dark respiration. As Glu and Gln appeared to be labelled by 13CO2, presumably their synthesis from 2-oxoglutarate molecules formed by the TCA cycle (by an isocitrate dehydrogenase) was favoured at the expense of succinate synthesis.

Nevertheless, under all experimental conditions (columns), Glu C-atom positions appeared to be less 13C enriched (up to 30%13C) than those in Mal (up to 66%13C), showing that Mal molecules formed via PEPc activity are not fully committed to Glu production, and/or that 13C-depleted, unlabelled molecules contribute to Glu production.

13C enrichment in key metabolites during a light : dark cycle

Figure 4 represents the relative 13C enrichment in key metabolites (C-3 citrate and average 13C enrichment in Ala, Asp, Glu, Gln and Mal) at three sampling times: (1) after 6 h light with 13CO2; (2) after 6 h light with 13CO2 and 30 min dark; and (3) after 6 h light with 13CO2 followed by 4 h darkness and 6 h light with CO2 at natural 13C abundance. These experiments were carried out under normal (white bars, Fig. 4) and high (black bars, Fig. 4) photorespiratory conditions; however, similar trends for each metabolite were observed at both CO2 levels. The 13C enrichment is relative to the C-5 atom position in glucose (Glc) in order to normalize the data by the photosynthetic isotopic input. The C-5 atom enrichment in Glc was chosen here as a reliable Glc peak for 13C measurements as it is quite separated from other signals in NMR spectra (carbohydrate region of chemical shifts) and, as such, minimizes overlap as a result of 13C–13C interactions within carbohydrates.

image

Figure 4.  Relative 13C enrichment detected by 13C-NMR after 6 h of 13CO2 labelling (column 1), 6 h of 13CO2 labelling and 30 min in darkness (column 2) and 6 h of 13CO2 labelling followed by 4 h in darkness and 6 h light in natural CO2 (1.1%13C) (column 3) in Brassica napus leaves. The CO2 mole fraction was 100 μmol mol−1 CO2 (filled bars) or 400 μmol mol−1 CO2 (open bars), and experiments were performed at 21% O2, 400 μmol m−2 s−1 photosynthetically active radiation (PAR) and 21°C. Ala, alanine; Asp, aspartate; Cit, citrate; Gln, glutamine; Glu, glutamate; Mal, malate. The values are the sum of 13C enrichments in all C-atom positions of the molecule of interest, relative to the 13C enrichment value in the C-5 atom position in Glc. Standard errors integrate the noise of each scan.

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Ala and Asp showed a quite low relative 13C enrichment after 6 h of 13CO2 labelling (sampling time 1), and then increased sharply in the dark (sampling time 2), thus indicating an increased commitment of 13C substrates to Asp and Ala synthesis in the dark. 13C-Ala mostly disappeared at sampling time 3, suggesting either an isotopic dilution by 12CO2 or a consumption of 13C-Ala in the light. In the present experiment, isotopic dilution was probably not sufficient to account for the lower 13C content in Ala because of the low 13C content obtained just after 6 h labelling with 13CO2 (sampling time 1). The same pattern occurred for Asp. We therefore conclude that, regardless of the photorespiratory conditions, Asp and Ala are synthesized both in the light and dark, and the simultaneous contribution of current assimilates and stored carbon dampens their isotopic variations in the light.

Citrate, Gln, Glu and Mal showed very similar patterns, with an increasing 13C enrichment from sampling time 1 to 3 (Fig. 4). This similar behaviour indicates a plausible common synthetic pathway. The very large increase in 13C enrichment on 12CO2 labelling (c. 10-fold relative to sampling time 1) was in contrast with that observed after 30 min in the dark (less than two-fold), thereby strongly suggesting that the synthesis of Glu that occurred in the light partly originated from 13C-enriched carbon from the previous light period. However, it may be argued that the increase in the relative 13C amount in citrate, Gln, Glu and Mal under 12CO2 labelling (Fig. 4) could be a result of the decrease in the amount of 13C in Glc C-5 (isotopic dilution by 12C). Nevertheless, the amount of 13C in Glc C-5 did not change from 6 h light + 30 min darkness to 6 h light + 4 h darkness + 6 h light, remaining close to 1.1% (Fig. 3). In other words, there was a clear increase in the average 13C amount in citrate, Gln, Glu and Mal in the light under 12CO2. Accordingly, the consumption of recent carbon that has just been fixed to produce Glu is small in the light: the percentages of recent carbon in Glu were 0% and 56% only after 1 h and 12 h labelling, respectively (calculated from Fig. 3). In other words, the turnover rate of carbon atoms in Glu in the light is as low as 0.046 mol mol−1 h−1. Such a pattern of 13C variations on labelling was not sensitive to photorespiratory conditions, as evidenced by very similar 13C enrichment variations under 100μmol mol−1 CO2 (lower plots, Fig. 4).

13C–15N spin–spin interactions

The 13C–15N spin–spin interaction observed by NMR between 13C and 15N nuclei is shown in Fig. 5. The interaction was quantified using either 13C-NMR (JCN, a) or 15N-NMR (JNC, b). The corresponding interpretation of the 13C chemical shift region associated with Glu is shown in Fig. 6. The average amount of 13C molecules that were also 15N labelled was nearly always < 60%. Quite surprisingly, the amount of 15N molecules that were also 13C labelled differed clearly, except for Gly and Ser, from that of the 13C molecules that were also 15N labelled. In other words, the populations of 15N and 13C molecules were dissimilar, with few common individuals.

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Figure 5. 13C–15N spin–spin interaction detected by 13C-NMR and 15N-NMR after 6 h of labelling with 13CO2 and 15NH415NO3 in Brassica napus leaves. The labelling was performed under either 100 μmol mol−1 CO2 (filled bars) or 400 μmol mol−1 CO2 (open bars), 21% O2, 400 μmol m−2 s−1 photosynthetically active radiation (PAR) and 21°C. JNC (a) and JCN (b) represent the proportion of 15N molecules interacting with 13C and the proportion of 13C molecules interacting with 15N, respectively. Asp, aspartate; Ala, alanine; Glu, glutamate; Gln, glutamine; Gly, glycine; Ser, serine.

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Figure 6.  Interpretation of the 13C chemical shift region between 54.25 and 56.25 ppm [C-2 atom position of glutamine (Gln)]. Spectra were obtained from Brassica napus leaves after 1, 6 or 12 h of 13CO2 labelling under 100 μmol mol−1 CO2 (a) or 400 μmol mol−1 CO2 (b). Experiments were performed at 21% O2, 400 μmol m−2 s−1 photosynthetically active radiation (PAR) and 21°C. Dichotomous schemes indicate the decomposition of 13C signals as a result of 13C–13C and 15N–13C interactions. After 1 h of labelling, no 13C signal was detected in Gln (lower spectra). Each graduation corresponds to a split of 25 Hz.

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Except for Gly and Ser, all 15N-Asp, 15N-Ala, 15N-Glu and 15N-Gln were not labelled with 13C, clearly showing that most of the recent 15N-amino groups were not fixed onto recent, 13C-enriched carbon skeletons. However, up to 60% of the 13C-oxoglutarate, 13C-pyruvate and 13C-oxaloacetate molecules formed in the light were fixed onto recent 15N-amino groups. This discrepancy indicates that 13C assimilation into amino acids is not tightly coupled to 15N assimilation, probably because of the involvement of stored 12C-carbon skeletons. The contrasting pattern in Gly and Ser was certainly related to their extremely large turnover associated with photorespiration, so that up to 50% of 15N-Gly and 15N-Ser molecules were also 13C labelled. This high turnover (Canvin, 1979) would explain the absence of differences observed between natural and high photorespiratory conditions in the present study.

Overall commitment of the 13C and 15N label into Glu

The relative 13C and 15N amounts in Glu (that is, the 13C and 15N isotopic signal divided by the total assimilated 13C and total absorbed 15N, respectively) were plotted against each other in Fig. 2, in which the data correspond to labelling with 13CO2 for 1, 6 and 12 h. Although the relative 13C amount increased with the amount of fixed CO2, the relative 15N amount decreased progressively as a result of the accumulation of 15N-ammonium nitrate in the leaf by the continuous absorption through the transpiration stream. At 400 μmol mol−1 CO2, the relative 13C amount nevertheless increased less between 6 and 12 h than between 1 and 6 h, that is, started to saturate. This presaturation was also visible in Fig. 3, as the specific 13C enrichment in Glu C-2 was slightly lower after 12 h than after 6 h at 400 μmol mol−1 CO2. Under high photorespiratory conditions (100 μmol mol−1 CO2), such a presaturation effect virtually disappeared, with a clear increase in the relative 13C amount between 6 and 12 h (Fig. 2) and a larger specific 13C enrichment in Glu C-2 at 12 h compared with 6 h (Fig. 2). In other words, there seemed to be a larger turnover of Glu under high photorespiration rates.

Discussion

  1. Top of page
  2. Summary
  3. Introduction
  4. Materials and Methods
  5. Results
  6. Discussion
  7. Acknowledgements
  8. References
  9. Supporting Information

Although being the cornerstone for nitrogen assimilation by plants, there remains much uncertainty about the origin of carbon skeletons required for Glu synthesis in illuminated leaves. Presumably, the carbon source for Glu production in the light comes from both newly synthesized (via photosynthates through the TCA cycle) and remobilized (derived from metabolites produced during the night) 2-oxoglutarate molecules. However, there is currently no corresponding assessment of the relative importance of such carbon sources. In this study, we took advantage of isotopic techniques and labelling with 13CO2 and 15N-ammonium nitrate, and carried out analyses with 13C- and 15N-NMR.

The TCA cycle in the light

The isotopic enrichment pattern of respiratory intermediates after 13CO2 feeding indicated that the TCA cycle was probably involved in Glu production in the light, as evidenced by the 13C enrichment in Glu and Gln (Fig. 3). The 13C enrichment pattern in citrate, Glu and Gln appeared to be quite similar (Fig. 4), suggesting that Glu and Gln are major carbon sinks for newly assimilated 13C atoms committed to the TCA cycle. Importantly, succinate was always poorly, if at all, labelled, strongly suggesting that the 13C label ([13C]-2-oxoglutarate) is directed to Glu/Gln synthesis at the expense of succinate. Such a conclusion is consistent with published biochemical data. Namely, 2-oxoglutarate dehydrogenation to succinyl-CoA is believed to be (1) impeded by the competition for Coenzyme A and E3 enzymatic subunits between pyruvate dehydrogenase and 2-oxoglutarate dehydrogenase (Dry & Wiskich, 1987; Budde et al., 1991; Millar & Kunst, 1999), and (2) inhibited by large NADH/NAD ratios within the mitochondrial matrix (Igamberdiev & Gärdestrom, 2003). That is, the production of succinate is quite low in the light.

Within such a framework, the involvement of PEPc was critical for regenerating oxaloacetate and supplementing citrate synthesis (anaplerotic role). In fact, Mal appeared to be labelled in all C-atom positions, and the 13C enrichment in C-4 covaried with the C-atom positions of Glu (the path followed by atoms is given in Fig. S3, see Supporting Information). Although leaf fumarate content was low (not detected with 13C-NMR), an equilibrium between Mal and fumarate probably occurred, thereby explaining the very similar 13C enrichment pattern in C-1 and C-4 atom positions in Mal (Fig. 3). The in vivo involvement of PEPc, which is activated in the light by phosphorylation (Duff & Chollet, 1995), has been further acknowledged by other metabolic studies that used isotopes at natural abundance (reviewed in Tcherkez & Hodges, 2008).

Our results thus indicate that oxaloacetate molecules produced by PEPc are committed to both Mal and citrate production, and that the TCA cycle does not appear to operate like a proper cycle but, rather, involves two opposite branches fed by PEPc. Such a scenario is in agreement with previous results obtained on cocklebur (Xanthium strumarium) leaves, in which Glu and Mal were more 13C labelled by 13C-2-pyruvate than was succinate (Tcherkez et al., 2008). Furthermore, 13CO2 labelling followed by fluxomics calculations (flux coefficients) in X. strumarium leaves have provided evidence that oxaloacetate molecules have two competitive fates in the light, that is, fumarate/Mal synthesis (reversed left branch of the cycle) and Glu synthesis (right branch of the cycle) (Tcherkez et al., 2009).

Notwithstanding the above observations, the 13C flux into the TCA cycle to Glu (right-hand side of the cycle) was rather small, on the order of 25 mmol 13C-Glu per mole of assimilated 13CO2 (Fig. 2), that is, 2.5% × 14 = 0.35 μmol m−2 s−1. This value is similar (same order of magnitude) to that of day respiration measured by gas exchange on several C3 plants (Atkin et al., 2000). Such a value also matches the in vitro citrate synthase activity measured on leaf extracts (illuminated leaves) of c. 0.3 μmol m−2 s−1, whereas that of PEPc is as large as c. 5 μmol m−2 s−1 in Nicotiana tabacum (Scheible et al., 2000). We therefore conclude that the restriction of day respiration and of the TCA cycle by light (see the references already discussed in the Introduction) is sufficient to provide the necessary 13C flux to feed Glu synthesis.

The contribution of stored carbon as a source for Glu synthesis

The production of Glu was further fed by the remobilization of stored molecules synthesized from previously assimilated carbon (‘old’ C atoms). In fact, the synthesis of 13C-Glu in the light was low compared with that observed in the subsequent dark period and in the next light period under ordinary CO2 (natural 13C abundance) (Fig. 4). This was also observed for citrate, Gln and Mal (Fig. 4). Such weak contributions of recent 13C atoms to Glu, Gln and citrate synthesis in the light were certainly not caused by the isotopic dilution of label 13CO2 by 12CO2 from photorespiratory and day respiratory decarboxylations. First, the photorespiratory intermediates Ser and Gly are rapidly labelled (Canvin, 1979), and this leads to photorespired CO2 becoming totally labelled. Second, the decarboxylation rate associated with day respiration is usually close to 0.5 μmol m−2 s−1 (Atkin et al., 2006), and therefore accounts for only 3% and 12% of the net photosynthetic CO2 exchange flux under 400 and 100 μmol mol−1 CO2, respectively. In addition, the proportion of refixed CO2 with respect to total decarboxylated CO2 is estimated to be close to 20% (Gerbaud & Andre, 1987; Tcherkez et al., 2005), thereby leading to a rather small isotopic dilution of fixed 13CO2 between 0.6 and 2%.

Other isotopic studies have also suggested that the production of respiratory CO2 in the light at the leaf (Parnik et al., 2002) or mesocosm (Schnyder et al., 2003) level relies on stored carbon up to a contribution of c. 50%. Accordingly, Huege et al. (2007) have shown that shifting from a 13CO2 atmosphere to a 12CO2 atmosphere in the light for 2 h is associated with virtually no decrease in the 13C content in Glu and succinate, thereby indicating that the role of stored carbon is critical. Furthermore, Mal and fumarate exhibited 13C half-times of more than 15 and 24 h, respectively, suggesting a large buffering pool of organic acids.

Our study shows that Asp and Ala are 13C depleted after a 6 h exposure to ordinary CO2 (following 6 h 13CO2 and 30 min in the dark), in clear contrast with Mal and Glu (Fig. 3). In addition, we obtained a rather low 13C enrichment in Asp after 6 h with 13CO2 in the light (Fig. 4, first bar), thereby showing that the production of Asp remains low in the light. It is therefore plausible that, in addition to the natural metabolic turnover (production from current photosynthetically fixed carbon), Asp and Ala are consumed and contribute to feeding oxaloacetate (Mal), pyruvate and Glu production in the light, respectively. It has already been noted that Asp levels are higher in Arabidopsis leaves during the night (Lam et al., 1995), such that, during the light period, it is believed to be either exported to sink tissues or used for Glu production from 2-oxoglutarate via the Asp to Glu transamination equilibrium. This view would also agree with the natural isotopic abundance in 15N: as the Asp to Glu transamination fractionates against 15N by 1.7‰ (Macko et al., 1986), a naturally 15N-enriched Asp leaf pool is expected, and this is indeed the case (Tcherkez & Hodges, 2008). Thus, a part of Glu in the light plausibly originates from Asp transamination, with Asp also synthesized in the light but at a lower rate than its consumption rate (‘dynamic equilibrium’).The contribution of stored carbon can explain why 15N and 13C labelling are dissimilar (Fig. 5). Indeed, although virtually no 15N-Glu or 15N-Gln molecules were also 13C labelled, up to 40% of 13C-Gln molecules were also 15N labelled. Similarly, when the relative isotopic 13C and 15N commitments in Glu were plotted against each other, there was a visible 15N enrichment at zero 13C enrichment (Fig. 2). In other words, within the first hour of labelling, Gln and Glu were hardly 13C labelled, but appeared to be 15N labelled by nearly 5%, and. accordingly, the 15N-NMR signal was rapidly detected under both ordinary and high photorespiratory conditions (Figs S1 and S2, see Supporting Information). This strongly suggests that Glu and Gln synthesis mainly involves carbon remobilization, but captures recently assimilated nitrogen. On the other hand, a substantial amount of newly synthesized, 13C-enriched 2-oxoglutarate is committed to nitrogen assimilation into Glu and Gln. Consistently, when leaves were transferred from a 13CO2 to a 12CO2 (natural 13C abundance) atmosphere, the 13C enrichment in citrate, Glu and Gln increased, undoubtedly showing the contribution of 13C intermediates stored during the night (Fig. 4).

Our scenario is also in agreement with the metabolic results obtained so far with mutants. First, tomato lines (Lycopersicon esculentum) with impaired fumarase activity showed no alterations in leaf Glu, pyro-Glu, succinate and glutarate contents in the light, whereas Mal and isocitrate tended to be larger (Nunes-Nesi et al., 2007). This indicates that fumarase is not essential for the regeneration of 2-oxoglutarate through the TCA cycle, and other carbon sources feed Glu production, such as stored organic acids and/or the anaplerotic PEPc activity, or the remaining fumarase activity is sufficient not to alter the carbon flow to nitrogen assimilation. Second, tomato lines with a reduced aconitase activity exhibited unchanged Glu and Gln levels when compared with wild-type plants in the light (but were much lower in darkness), whereas aspartate, succinate and fumarate contents were significantly lower and (iso)citrate accumulated (Carrari et al., 2003). Clearly then, the production of 2-oxoglutarate and Glu is independent of the variations in other TCA intermediates, indicating that the recycling of stored carbon skeletons to Glu occurs. Third, potato lines (Solanum tuberosum) with increased PEPc activity (2.7- to 4.7-fold) showed a larger Glu to Gln ratio, as well as pyruvate and 2-oxoglutarate contents (Rademacher et al., 2002), suggesting that the anaplerotic input by PEPc is essential for providing 2-oxoglutarate and Glu. In addition, in tobacco lines (Nicotiana sylvestris) in which the nitrate reductase activity is increased, the Gln to Glu ratio has been found to be exacerbated, whereas the Glu level is unaffected, so that it is probable that the production of 2-oxoglutarate is not limiting (Gojon et al., 1998).

Taken as a whole, such a lack of correlation between TCA intermediates and Glu levels points to a significant role for night-stored carbon (such as organic acids) in nitrogen assimilation and Glu synthesis in the light. This view is fully consistent with fluxomics calculations made in X. strumarium leaves, in which citrate synthase activity appears to be limiting, so that Glu synthesis in the light is associated with citrate remobilization (Tcherkez et al., 2009). In fact, the amount of organic acids, such as citrate, is likely to be sufficient to sustain Glu synthesis in the light. According to the fluxes found by Tcherkez et al. (2005), the rate associated with day respiratory CO2 evolution is c. 0.5 μmol m−2 s−1, whereas that associated with TCA activity is c. 0.05 μmol m−2 s−1. Therefore, Glu production is within the 0.05–0.5 μmol m−2 s−1 range (a value of 0.35 μmol m−2 s−1 was estimated here, see above). This would represent a carbon skeleton requirement of c. 0.35 × 6 × 3600 = 7.5 mmol m−2 during a 6 h illumination period. Under our experimental conditions, the Mal content was close to 7 mmol m−2 after 6 h illumination (data not shown), which is adequate to complete a light period of 12 h. Although we recognize that the Mal content may vary during the night (Gerhardt & Heldt, 1984), it thus seems that the sum of Mal and citrate contents available at the very beginning of the light period is sufficient to provide nearly all the 2-oxoglutarate molecules required for Glu production.

The overall commitment of the 13C label into Glu was c. 25–30 mmol per mole of net assimilated CO2, regardless of the photorespiratory conditions (Fig. 2), that is, on the order of 3%. Previous 14C-labelling experiments gave similar results in Arabidopsis thaliana and spinach (Spinacia oleracea), with values of c. 6 and 3%, respectively (Lawyer et al., 1981; Carrari et al., 2003). Photorespiration had a stimulating effect on such a 13C commitment, that is, on 13C-Glu neosynthesis (Figs 2,3). Accordingly, labelling experiments with 14CO2 and 13C-pyruvate have shown that Glu synthesis is promoted under photorespiratory conditions (Lawyer et al., 1981; Tcherkez et al., 2008): Glu and/or Gln represented a larger amount of isotopic label and a higher isotopic specific activity after labelling in ordinary conditions relative to nonphotorespiratory conditions. Still, the contribution of stored carbon to Glu synthesis was apparent (i.e. weakly sensitive to photorespiration), with the clear production of 13C-Glu when the experimental conditions turned to 12CO2 conditions, regardless of the CO2 mole fraction (Fig. 4).

Rationale and perspectives

Interactions between current carbon assimilation, day respiration and photorespiration are quite complex because photorespiration, PEPc activity and the TCA cycle are associated with amino acid metabolism (Gly, Ser, Asp, Glu, Gln). Although Glu and 2-oxoglutarate are involved in a cycle (the photorespiratory Glu recovery cycle), and may thus be assumed to have stationary levels, the shift from steady ordinary (380 μmol mol−1 CO2; 21% O2) to large values of both the oxygenation to carboxylation ratio (vo/vc) and nitrogen assimilation into Glu require net 2-oxoglutarate synthesis. Our results indicate that the remobilization of night-stored molecules, such as organic acids (e.g. citrate, see Scheible et al., 2000 and Niedziela et al., 1993) or amino acids (e.g. Asp and Ala), plays a major role in feeding 2-oxoglutarate synthesis for nitrogen assimilation. Indeed, our results show that the natural day : night cycle is critical for nitrogen assimilation, as intermediates produced in the dark are required during the subsequent light period for nitrogen reduction and assimilation in leaves (Canvin & Atkins, 1974; Pilgrim et al., 1993; Scheible et al., 2000; Lillo et al., 2001).

Within such a framework, the well-recognized relationship between nitrogen assimilation and maintenance respiration in darkness (for a review, see Thornley & Cannell, 2000) comes as no surprise. Similarly, although the nitrogen content in leaves may vary strongly between species (up to two-fold), leaf night respiration per nitrogen content unit remains constant (at c. 10 nmol CO2 nmol−1 N s−1), again suggesting that nitrogen assimilation correlates with night respiratory metabolism (Poorter et al., 1990).

We nevertheless recognize that such a metabolic compromise along the day : night cycle may depend on environmental conditions, such as temperature or light, that influence respiration, which may, in turn, change nitrate reduction and assimilation rates. For example, during growth under a CO2-enriched atmosphere and nitrogen-limited conditions, there is a marked decrease in nitrate reductase activity and amino acid concentration (Geiger et al., 1999), as well as plant nitrogen concentration (percentage of nitrogen), with both maintenance and basal dark-adapted respiration decreasing accordingly (Gifford & Bayer, 1995). That said, the whole-plant respiratory cost R : P is weakly, if at all, affected by CO2 conditions (Gifford & Bayer, 1995; Albrizio & Steduto, 2003), indicating that the control on the commitment of assimilates to respiration is critical for nitrogen assimilation. Furthermore, some evidence has been provided that nitrogen uptake and assimilation in roots are dependent on the input of carbon assimilates that are, in turn, used as respiratory substrates (Gojon et al., 1991). Therefore, the control of nitrogen assimilation by respiration at the whole-plant scale is further complicated by allocation patterns and shoot : root ratios, and, consequently, there remains much uncertainty as to whether our results are still valid on a long-term basis when environmental conditions vary; this will be addressed in a subsequent study.

Acknowledgements

  1. Top of page
  2. Summary
  3. Introduction
  4. Materials and Methods
  5. Results
  6. Discussion
  7. Acknowledgements
  8. References
  9. Supporting Information

The authors are very grateful to Dr Nathalie Nesi for providing rape seeds. The authors wish to thank the Agence Nationale de la Recherche and the IFR87 for financial support through a Jeunes Chercheur project (under contract 08-330055) and a Transversal Project, respectively. P.G. was financed by a PhD Grant from the French Ministère de l’Éducation Nationale. G.T. and M.H. wish to thank Professor Gabriel Cornic for his strong support for this work.

References

  1. Top of page
  2. Summary
  3. Introduction
  4. Materials and Methods
  5. Results
  6. Discussion
  7. Acknowledgements
  8. References
  9. Supporting Information
  • Albrizio R, Steduto P. 2003. Photosynthesis, respiration and conservative carbon use efficiency of four field grown crops. Agricultural and Forest Meteorology 116: 1936.
  • Atkin OK, Millar AH, Gärdestrom P, Day DA. 2000. Photosynthesis, carbohydrate metabolism and respiration in leaves of higher plants. In: LeegoodRC, SharkeyTD, von CaemmererS, eds. Photosynthesis, physiology and metabolism. London, UK: Kluwer Academic Publishers, 203220.
  • Atkin OK, Scheurwater I, Pons TL. 2006. High thermal acclimation potential of both photosynthesis and respiration in two lowland Plantago species in contrast to an alpine congeneric. Global Change Biology 12: 500515.
  • Aubert S, Gout E, Bligny R, MartyMazars D, Barrieu F, Alabouvette J, Marty F, Douce R. 1996. Ultrastructural and biochemical characterization of autophagy in higher plant cells subjected to carbon deprivation: control by the supply of mitochondria with respiratory substrates. Journal of Cell Biology 133: 12511263.
  • Bauer A, Urquhart AA, Joy KW. 1977. Amino acid metabolism of pea leaves: diurnal changes and amino acid synthesis from 15N-nitrate. Plant Physiology 59: 915919.
  • Budde RJA, Fang TK, Randall DD, Miernyk JA. 1991. Acetyl-coenzyme-a can regulate activity of the mitochondrial pyruvate-dehydrogenase complex in-situ. Plant physiology 95: 131136.
  • Canvin DT. 1979. Photorespiration: comparison between C3 and C4 plants. In: GibbsM, LatzckoE, eds. Photosynthesis II. Photosynthetic carbon metabolism and related processes. Springer-Verlag, Berlin: 368396.
  • Canvin DT, Atkins CA. 1974. Nitrate, nitrite and ammonia assimilation by leaves – effect of light, carbon-dioxide and oxygen. Planta 116: 207224.
  • Carrari F, Nunes-Nesi A, Gibon Y, Lytovchenko A, Loureiro ME, Fernie AR. 2003. Reduced expression of aconitase results in an enhanced rate of photosynthesis and marked shifts in carbon partitioning in illuminated leaves of wild species tomato. Plant Physiology 133: 13221335.
  • Delhon P, Gojon A, Tillard P, Passama L. 1995. Diurnal regulation of NO3 uptake in soybean plants 1. Changes in NO3 influx, efflux, and N utilization in the plant during the day–night cycle. Journal of Experimental Botany 46: 15851594.
  • Dry IB, Wiskich JT. 1987. 2-Oxoglutarate dehydrogenase and pyruvate-dehydrogenase activities in plant-mitochondria – interaction via a common coenzyme-a pool. Archives of Biochemistry and Biophysics 257: 9299.
  • Duff SMG, Chollet R. 1995. In-vivo regulation of wheat-leaf phosphoenolpyruvate carboxylase by reversible phosphorylation. Plant Physiology 107: 775782.
  • Forde BG, Lea PJ. 2007. Glutamate in plants: metabolism, regulation and signalling. Journal of Experimental Botany 58: 23392358.
  • Geiger M, Haake V, Ludewig F, Sonnewald U, Stitt M. 1999. The nitrate and ammonium nitrate supply have a major influence on the response of photosynthesis, carbon metabolism, nitrogen metabolism and growth to elevated carbon dioxide in tobacco. Plant, Cell & Environment 22: 11771199.
  • Gerbaud A, Andre M. 1987. An evaluation of the recycling in measurements of photorespiration. Plant Physiology 83: 933937.
  • Gerhardt R, Heldt HW. 1984. Measurement of subcellular metabolite levels in leaves by fractionation of freeze-stopped material in nonaqueous media. Plant Physiology 75: 542547.
  • Gifford EM, Bayer DE. 1995. Developmental anatomy of Cyperus esculentus (yellow nutsedge). International Journal of Plant Sciences 156: 622629.
  • Gojon A, Passama L, Robin P. 1986. Root contribution to nitrate reduction in barley seedlings (Hordeum vulgare-L). Plant and Soil 91: 339342.
  • Gojon A, Bussi C, Grignon C, Salsac L. 1991. Distribution of NO3 reduction between roots and shoots of peach tree seedlings as affected by NO3 uptake rate. Physiologia Plantarum 82: 505512.
  • Gojon A, Dapoigny L, Lejay L, Tillard P, Rufty TW. 1998. Effects of genetic modification of nitrate reductase expression on (NO3)-N-15 uptake and reduction in Nicotiana plants. Plant, Cell & Environment 21: 4353.
  • Gout E, Bligny R, Pascal N, Douce R. 1993. 13C Nuclear Magnetic Resonance studies of malate and citrate synthesis and compartmentation in higher plant cells. The Journal of Biological Chemistry 268: 39863992.
  • Guo S, Schinner K, Sattelmacher B, Hansen UP. 2005. Different apparent CO2 compensation points in nitrate- and ammonium-grown Phaseolus vulgaris and the relationship to non-photorespiratory CO2 evolution. Physiologia Plantarum 123: 288301.
  • Hanning I, Heldt HW. 1993. On the function of mitochondrial metabolism during photosynthesis in spinach (Spinacia oleracea L) leaves – partitioning between respiration and export of redox equivalents and precursors for nitrate assimilation products. Plant Physiology 103: 11471154.
  • Huege J, Sulpice R, Gibon Y, Lisec J, Koehl K, Kopka J. 2007. GC-EI-TOF-MS analysis of in vivo carbon-partitioning into soluble metabolite pools of higher plants by monitoring isotope dilution after (CO2)-C-13 labelling. Phytochemistry 68: 22582272.
  • Huppe HC, Turpin DH. 1994. Integration of carbon and nitrogen metabolism in plant and algal cells. Annual Review of Plant Physiology and Plant Molecular Biology 45: 577607.
  • Igamberdiev AU, Gärdestrom P. 2003. Regulation of NAD- and NADP-dependent isocitrate dehydrogenases by reduction levels of pyridine nucleotides in mitochondria and cytosol of pea leaves. Biochimica et Biophysica Acta-Bioenergetics 1606: 117125.
  • Kruse A, Fieuw S, Heineke D, Muller-Rober B. 1998. Antisense inhibition of cytosolic NADP-dependent isocitrate dehydrogenase in transgenic potato plants. Planta 205: 8291.
  • Lam HM, Coschigano K, Schultz C, Melooliveira R, Tjaden G, Oliveira I, Ngai N, Hsieh MH, Coruzzi G. 1995. Use of Arabidopsis mutants and genes to study amide amino-acid biosynthesis. Plant Cell 7: 887898.
  • Lawlor DW. 2002. Carbon and nitrogen assimilation in relation to yield: mechanisms are the key to understanding production systems. Journal of Experimental Botany 53: 773787.
  • Lawyer AL, Cornwell KL, Larsen PO, Bassham JA. 1981. Effects of carbon dioxide and oxygen on the regulation of photosynthetic carbon metabolism by ammonia in spinach mesophyll cells. Plant Physiology 68: 12311236.
  • Lemaitre T, Urbanczyk-Wochniak E, Flesch V, Bismuth E, Fernie AR, Hodges M. 2007. NAD-dependent isocitrate dehydrogenase mutants of Arabidopsis suggest the enzyme is not limiting for nitrogen assimilation. Plant Physiology 144: 15461558.
  • Lillo C, Meyer C, Ruoff P. 2001. The nitrate reductase circadian system. The central clock dogma contra multiple oscillatory feedback loops. Plant Physiology 125: 15541557.
  • Macko SA, Estep MLF, Engel MH, Hare PE. 1986. Kinetic fractionation of stable nitrogen isotopes during amino-acid transamination. Geochimica et Cosmochimica Acta 50: 21432146.
  • Maillard P, Deleens E, Daudet FA, Lacointe A, Frossard JS. 1994. Carbon and nitrogen partitioning in walnut seedlings during the acquisition of autotrophy through simultaneous (CO2)-C-13 and (NO3)-N-15 long-term labeling. Journal of Experimental Botany 45: 203210.
  • Millar AA, Kunst L. 1999. The natural genetic variation of the fatty-acyl composition of seed oils in different ecotypes of Arabidopsis thaliana. Phytochemistry 52: 10291033.
  • Niedziela CE, Nelson PV, Peet MM, Jackson WA. 1993. Diurnal malate and citrate fluctuations as related to nitrate and potassium concentrations in tomato leaves. Journal of Plant Nutrition 16: 165175.
  • Nunes-Nesi A, Carrari F, Gibon Y, Sulpice R, Lytovchenko A, Fisahn J, Graham J, Ratcliffe RG, Sweetlove LJ, Fernie AR. 2007. Deficiency of mitochondrial fumarase activity in tomato plants impairs photosynthesis via an effect on stomatal function. Plant Journal 50: 10931106.
  • Parnik TR, Voronin PY, Ivanova HN, Keerberg OF. 2002. Respiratory CO2 fIuxes in photosynthesizing leaves of C-3 species varying in rates of starch synthesis. Russian Journal of Plant Physiology 49: 729735.
  • Pilgrim ML, Caspar T, Quail PH, McClung CR. 1993. Circadian and light regulated expression of nitrate reductase in Arabidopsis. Plant Molecular Biology 23: 349364.
  • Poorter H, Remkes C, Lambers H. 1990. Carbon and nitrogen economy of 24 wild species differing in relative growth rate. Plant Physiology 94: 621627.
  • Rademacher T, Hausler RE, Hirsch HJ, Zhang L, Lipka V, Weier D, Kreuzaler F, Peterhansel C. 2002. An engineered phosphoenolpyruvate carboxylase redirects carbon and nitrogen flow in transgenic potato plants. Plant Journal 32: 2539.
  • Radin JW. 1977. Contribution of root-system to nitrate assimilation in whole cotton plants. Australian Journal of Plant Physiology 4: 811819.
  • Radin JW. 1978. Physiological basis for division of nitrate assimilation between roots and leaves. Plant Science Letters 13: 2125.
  • Saeed AI, Sharov V, White J, Li J, Liang W, Bhagabati N, Braisted J, Klapa M, Currier T, Thiagarajan M et al. 2003. TM4: a free, open-source system for microarray data management and analysis. BioTechniques 34: 374378.
  • Scheible WR, Krapp A, Stitt M. 2000. Reciprocal diurnal changes of phosphoenolpyruvate carboxylase expression and cytosolic pyruvate kinase, citrate synthase and NADP-isocitrate dehydrogenase expression regulate organic acid metabolism during nitrate assimilation in tobacco leaves. Plant, Cell & Environment 23: 11551167.
  • Schnyder H, Schaufele R, Lotscher M, Gebbing T. 2003. Disentangling CO2 fluxes: direct measurements of mesocosm-scale natural abundance (CO2)-C-13/(CO2)-C-12 gas exchange, C-13 discrimination, and labelling of CO2 exchange flux components in controlled environments. Plant, Cell & Environment 26: 18631874.
  • Stitt M, Muller C, Matt P, Gibon Y, Carillo P, Morcuende R, Scheible WR, Krapp A. 2002. Steps towards an integrated view of nitrogen metabolism. Journal of Experimental Botany 53: 959970.
  • Tcherkez G, Hodges M. 2008. How stable isotopes may help to elucidate primary nitrogen metabolism and its interaction with (photo)respiration in C-3 leaves. Journal of Experimental Botany 59: 16851693.
  • Tcherkez G, Cornic G, Bligny R, Gout E, Ghashghaie J. 2005. In vivo respiratory metabolism of illuminated leaves. Plant Physiology 138: 15961606.
  • Tcherkez G, Ghashghaie J, Griffiths H. 2007. Methods for improving the visualization and deconvolution of isotopic signals. Plant, Cell & Environment 30: 887891.
  • Tcherkez G, Bligny R, Gout E, Mahe A, Hodges M, Cornic G. 2008. Respiratory metabolism of illuminated leaves depends on CO2 and O2 conditions. Proceedings of the National Academy of Sciences, USA 105: 797802.
  • Tcherkez G, Mahe A, Gauthier P, Mauve C, Gout E, Bligny R, Cornic G, Hodges M. 2009. In folio respiratory fluxomics revealed by 13C-isotopic labeling and H/D isotope effects highlight the non-cyclic nature of the tricarboxylic acid “cycle” in illuminated leaves. Plant Physiology 151: 620630.
  • Thornley JHM, Cannell MGR. 2000. Modelling the components of plant respiration: representation and realism. Annals of Botany 85: 5567.
  • Vartanian N, Damerval C, de Vienne D. 1987. Drought-induced changes in protein patterns of Brassica napus var oleifera roots. Plant Physiology 84: 989992.