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First published online November 17, 2006; 10.1104/pp.106.091330

Plant Physiology 143:530-539 (2007)
© 2007 American Society of Plant Biologists

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SYSTEMS BIOLOGY, MOLECULAR BIOLOGY, AND GENE REGULATION

Tobacco Isoenzyme 1 of NAD(H)-Dependent Glutamate Dehydrogenase Catabolizes Glutamate in Vivo[OA]

Matthew Peter Purnell1 and José Ramon Botella*

Department of Botany, School of Integrative Biology, University of Queensland, Queensland 4072, Australia


   ABSTRACT
TOP
 ABSTRACT
RESULTS
 DISCUSSION
 CONCLUSION
 MATERIALS AND METHODS
 LITERATURE CITED
 
Glutamate (Glu) dehydrogenase (GDH, EC 1.4.1.2–1.4.1.4) catalyzes in vitro the reversible amination of 2-oxoglutarate to Glu. The in vivo direction(s) of the GDH reaction in higher plants and hence the role(s) of this enzyme is unclear, a situation confounded by the existence of isoenzymes comprised totally of either GDH beta- (isoenzyme 1) or {alpha}- (isoenzyme 7) subunits, as well as another five {alpha}-beta isoenzyme permutations. To clarify the in vivo direction of the reaction catalyzed by GDH isoenzyme 1, [15N]Glu was supplied to roots of two independent transgenic tobacco (Nicotiana tabacum) lines with increased isoenzyme 1 levels (S4-H and S49-H). The [15N]ammonium (NH4+) accumulation rate in these lines was elevated approximately 65% compared with a null segregant control line, indicating that isoenzyme 1 catabolizes Glu in roots. Leaf glutamine synthetase (GS) was inhibited with a GS-specific herbicide to quantify any contribution by GDH toward photorespiratory NH4+ reassimilation. Transgenic line S49-H did not show enhanced resistance to the herbicide, indicating that the large pool of isoenzyme 1 in S49-H leaves was unable to compensate for GS and suggesting that isoenzyme 1 does not assimilate NH4+ in vivo.

The intricate linkage of plant C and N metabolism requires constant balancing at the cellular level. Fundamentally, this involves orchestrating cellular levels of three metabolites: ammonium (NH4+), the N form assimilated into organic molecules; 2-oxogluturate (2-OG), the C skeleton to which NH4+ is most often joined; and Glu, the amino acid resulting from NH4+ and 2-OG condensation (Lancien and Gadal, 2000Go; Coruzzi and Zhou, 2001Go).

Glutamate dehydrogenase (GDH; EC 1.4.1.2) catalyzes a reaction incorporating these metabolites, and hence, this enzyme constitutes a crossroads of C and N metabolism. In vitro, the reversible, near-equilibrium reaction catalyzed by GDH is: 2-OG + NH4+ + NADH {leftrightarrow} Glu + NAD+ + water. Before 1974, GDH was thought to be the major portal for NH4+ assimilation in higher plants. However, in 1974, this function was attributed to the newly discovered Gln synthetase (GS)/GOGAT cycle (Lea and Miflin, 1974Go). Subsequently, many attempts have been made to redefine the role of GDH, but this has remained elusive and controversial because of difficulties in determining the in vivo direction of the GDH reaction. There is evidence that if GDH operates in the anabolic direction, it may reassimilate a small amount of photorespiratory NH4+ and/or help assimilate excessive exogenous NH4+. Conversely, if GDH operates in the catabolic direction, it may help alter cellular C to N ratios and/or fuel the TCA cycle under conditions of C deficit (Rhodes et al., 1989Go; Robinson et al., 1991Go, 1992Go; Aubert et al., 2001Go; Loulakakis and Roubelakis-Angelakis, 2001Go; Miflin and Habash, 2002Go; Dubois et al., 2003Go; Skopelitis et al., 2006Go).

The difficulty of determining the in vivo direction of the GDH reaction is exacerbated by the existence of seven GDH isoenzymes that result from the association of two subunit types ({alpha} and beta) in the GDH hexamer (i.e. beta6 [isoenzyme 1], beta5{alpha}1...beta1{alpha}5, {alpha}6 [isoenzyme 7]; Cammaerts and Jacobs, 1983Go; Loulakakis and Roubelakis-Angelakis, 1990Go; Magalhaes et al., 1990Go). The GDH isoenzyme profile of a particular organ can change during development (e.g. onset of leaf senescence; Cammaerts and Jacobs, 1985Go). Therefore, although the GDH homohexamers have near-identical kinetic properties in vitro (Loulakakis and Roubelakis-Angelakis, 1996Go), it is possible that they may catalyze opposite reactions in vivo and hence have different physiological roles. {alpha}- and beta-subunit cDNAs have been cloned from several species (summarized by Purnell et al., 2005Go), and gene expression studies to date have demonstrated generally that subunit gene expression is promoted by a low cellular C to N ratio (e.g. in C sinks, during darkness, and during C starvation of explants) and repressed by a high cellular C to N ratio (e.g. in C sources, during light, and during supply of Suc to explants; Melo-Oliveira et al., 1996Go; Turano et al., 1997Go; Ficarelli et al., 1999Go; Masclaux et al., 2000Go; Masclaux-Daubresse et al., 2002Go; Tercé-Laforgue et al., 2004bGo), agreeing with GDH activity data (Restivo et al., 1989Go; Robinson et al., 1992Go; Masclaux et al., 2000Go; Morkunas et al., 2000Go; Tercé-Laforgue et al., 2004bGo). In source leaves, both subunits are localized to mitochondria of phloem companion cells. In leaves with elevated NH4+ levels (e.g. senescing leaves), GDH protein is also detected in the cytosol of phloem companion cells (Paczek et al., 2002Go; Dubois et al., 2003Go; Tercé-Laforgue et al., 2004aGo; Kichey et al., 2005Go; Fontaine et al., 2006Go).

Metabolic studies with a maize (Zea mays) beta-subunit deficit mutant, which has a 10- to 15-fold decrease in root GDH activity (Magalhaes et al., 1990Go; Pryor, 1990Go), have provided the best evidence for the in vivo direction of the GDH isoenzyme 1 reaction. Assimilation of [15N]NH4+ supplied to roots was decreased 40% to 50% in the mutant, suggesting that GDH isoenzyme 1 assimilates NH4+ in roots (Magalhaes et al., 1990Go). However, the authors urged caution in making this conclusion, because the mutant had an approximately 20% smaller shoot-to-root ratio (due to a greater root mass), and in the presence of the potent GS inhibitor, Met sulfoximine (MSX), NH4+ assimilation was prevented. In another study, the large reduction in root GDH activity in the mutant was associated with a corresponding reduction in root [15N]Glu catabolism. However, [15N]Glu uptake by mutant roots was only 25% of that of wild-type roots, making direct comparison difficult (Stewart et al., 1995Go). Besides the results of Magalhaes et al. (1990)Go and Stewart et al. (1995)Go being apparently conflicting, the interpretation of these results is complicated by a lack of an isogenic control, making it unclear whether the altered morphology and [15N]Glu uptake rate of the maize GDH mutant are pleiotropic effects of the mutation or rather merely a function of plant variety.

Transgenic plants with modulated GDH {alpha}- or beta-subunit levels have recently been reported (Purnell et al., 2005Go; Fontaine et al., 2006Go). Purnell et al. (2005)Go generated transgenic tobacco (Nicotiana tabacum) lines with increased beta-subunit and isoenzyme 1 levels. Unlike the maize GDH beta-subunit mutant, the morphology of T2-generation transformants was not noticeably altered compared to isogenic null segregant (NS) control lines (Purnell et al., 2005Go). In this article, we have performed metabolic studies with these transgenic lines. Our results indicate that in roots, GDH isoenzyme 1 catabolizes Glu.


   RESULTS
TOP
 ABSTRACT
 RESULTS
DISCUSSION
 CONCLUSION
 MATERIALS AND METHODS
 LITERATURE CITED
 

Effect of Increasing GDH Isoenzyme 1 Content on Metabolism of Supplied [15N]Glu

We had previously generated multiple independent transgenic tobacco lines with increased GDH isoenzyme 1 levels (Purnell et al., 2005Go). To investigate the effect of a large GDH isoenzyme 1 pool on Glu metabolism, two of these lines (S4-H and S49-H) and an isogenic NS control (A63-NS) were supplied to [15N]Glu via the roots for 4 h. In addition to this treatment (henceforth the control treatment), the effect of inhibiting either GS or all pyridoxal-P-dependent enzymes (including aminotransferases and amino acid decarboxylases) was investigated by cosupplying either MSX (MSX treatment) or aminooxyacetate (AOA; AOA treatment), respectively. A fourth treatment consisted of simultaneously cosupplying both MSX and AOA.

The metabolism of [15N]Glu was investigated in roots and leaves by NMR spectroscopy. To maximize sensitivity, the 15N nucleus was detected indirectly using a spectral-editing pulse sequence, which detected only those 1H resonances arising from 15N-labeled compounds. After optimizing the detection of labeled NH4+, Glu, and Gln (Fig. 1 ), this method was approximately 2-fold more sensitive and 9 times faster than direct detection of the 15N nucleus.


Figure 1
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  		Figure 1. Edited NMR 1H spectra of reference compounds. Optimization of signals for 15N-labeled Gln amide (A; 11.4 mM), Glu (B; 9 mM), and NH4+ (C; 20 mM). D, Unedited spectrum. Spectra not to scale.

 
In A63-NS roots, the label was detected in NH4+, Glu, and the amide of Gln (Fig. 2 ). The 1H and 15N chemical shifts ({delta}) and the concentrations of these compounds are listed in Table I . An additional peak at {delta}1H = 7.69 ppm was attributed to [15N]{gamma}-aminobutyric acid (GABA) via two-dimensional heteronuclear single-quantum coherence (2D HSQC) NMR (inset Fig. 2), while another peak at {delta}1H = 8.08 ppm was not identified. The 2D spectrum indicated a {delta}15N = 106.64 ppm for this unidentified compound but did not reveal any 1H correlations (results not shown). Mesnard et al. (2000)Go, working with Nicotiana plumbaginifolia cell cultures, identified two 15N-labeled compounds with a similar {delta}15N that also had no 1H correlations in a 2D HSQC spectrum; namely, N-acetyl Glu (107.5 ppm) and N-acetyl Orn (107.4 ppm). It is possible that the unknown compound in this study is one of these compounds, although it should be noted that acquisition-parameter differences (e.g. pH and temperature) between this study and that of Mesnard et al. (2000)Go would have caused {delta}15N differences. Labeled NH4+ and GABA also accumulated in leaves of A63-NS (results not shown), but labeled Glu and Gln were below the level of detection. At 4 h, the concentration of [15N]NH4+ in leaves was 160 nmol g fresh weight–1, or 3.2% of the amount in roots. Although absolute quantification of [15N]GABA in roots and leaves was not conducted, [15N]GABA levels in A63-NS leaves were only 7.3% of the levels detected in roots.


Figure 2
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  		Figure 2. Metabolism of 15N[Glu] by roots of the control line A63-NS in the absence of enzyme inhibitors. Optimized detection for 15N-labeled Gln amide (A), Glu (B), and NH4+ (C). D, Unedited 1H spectrum. ?, Unidentified compound. Spectra not to scale. Inset shows assignment of resonances in a 2D HQSR-TOSCY spectrum of an extract from roots treated with MSX. 15N chemical shifts ({delta}) are indicated by the y axis, and {delta} of correlating protons by the x axis.

 

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  		Table I. Chemical shifts, concentrations, and accumulation rates of compounds detected in roots of the control line A63-NS 4 h after supply of 5 mM [15N]Glu

 
MSX treatment prevented accumulation of labeled Gln, Glu, and the unidentified compound in A63-NS roots (compare Fig. 3, A and B, and C and D ), indicating that the unidentified compound was produced via GS activity. [15N]GABA accumulation increased approximately 60% (P < 0.05) in MSX-treated roots compared with nontreated roots (compare Fig. 3, C and D), due probably to increased access of Glu decarboxylase (GAD; EC 4.1.1.15; Bouche and Fromm, 2004Go) to the [15N]Glu pool. Furthermore, a 2D spectrum of MSX-treated roots (Fig. 2, inset) revealed a small amount of [15N]Ala that was not detected in nontreated roots (results not shown). This [15N]Ala was formed probably by mitochondrial GABA transaminase (EC 2.6.1.19) transferring the amino group of [15N]GABA to pyruvate (Cauwenberghe and Shelp, 1999Go). No other labeled amino acids were detected, indicating that other aminotransferase activities may have been negligible.


Figure 3
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  		Figure 3. Effect of enzyme inhibitors on 15N[Glu] metabolism by A63-NS roots. A and B, Acquisition parameters set for optimal detection of labeled Gln amide arising from the control treatment (A) or from application of MSX (B). C to F, Acquisition parameters set for optimal detection of [15N]NH4+ arising from the control treatment (C), application of MSX (D), application of AOA (E), or application of MSX and AOA (F). ?, Unidentified compound. A and B to scale, and C to F to scale.

 
AOA treatment prevented accumulation of [15N]GABA in A63-NS roots and leaves (compare Fig. 3, D and E), indicating that all of the [15N]GABA detected in the control treatment was most probably derived from GAD activity. The combined MSX + AOA treatment prevented the accumulation of all labeled compounds except [15N]NH4+ in A63-NS roots and leaves (Fig. 3F), indicating that a substantial amount of [15N]NH4+ was liberated from 15N[Glu] independently of GS, GAD, or aminotransferase activity.

[15N]GABA content was determined for all genotypes 4 h after supply of the label, and in addition, A63-NS was also assayed 2 h after label supply. Because GAD activity increases with increasing cytosolic Glu concentrations (Scott-Taggart et al., 1999Go), in this study, GAD-produced [15N]GABA was a convenient marker of cytosolic [15N]Glu content. [15N]GABA content was similar for each genotype, and the [15N]GABA accumulation rate within A63-NS roots was linear (results not shown), indicating a similar and linear [15N]Glu uptake rate for all genotypes.

For each genotype, [15N]NH4+ accumulation in roots and leaves was followed over time in the absence or presence of MSX (Fig. 4 ). In the absence of MSX, the overexpressing lines S4-H and S49-H accumulated 68% and 62% more [15N]NH4+ in roots than the control line, A63-NS, respectively (Fig. 4A, 4 h; P = 0.0015). In leaves, differences between A63-NS and the transgenic lines were not significant (P > 0.05; Fig. 4C). Treatment with MSX resulted in a similar pattern of [15N]NH4+ accumulation in roots to that observed in the control treatment (Fig. 4B). [15N]NH4+ content was 28% and 26% higher in S4-H and S49-H, respectively, compared with A63-NS (Fig. 4B, 4 h; P = 0.0029). Again, no significant differences were observed in leaves between A63-NS and the transgenic lines (Fig. 4D; P > 0.05).


Figure 4
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  		Figure 4. Accumulation of 15N[NH4+] in roots and leaves of GDH beta-subunit overexpressers supplied with 15N[Glu]. Hydroponically grown plants were supplied 15N[Glu] via the roots without (control treatment; A and C) or with (B and D) MSX added to the medium. Accumulation of 15N[NH4+] over time was measured in roots (A and B) and leaves (C and D) of the control line, A63-NS ({blacksquare}), and GDH beta-subunit overexpressing lines, S49-H ({blacktriangleup}), and S4-H (bullet). Data are the means ± SE (n = 3). Error bars are not shown where smaller than the symbol and to improve clarity are sometimes shown in one direction only. For A and B, there was a significant difference (P ≤ 0.0043) among the lines at 2 and 4 h, as determined by a two-way ANOVA. For each time point, means not marked with the same letter were significantly different.

 

Ability of a GDH Overexpressing Line to Reassimilate Photorespiratory NH4+

To ascertain whether a large ectopic pool of GDH isoenzyme 1 in the leaves can reassimilate photorespiratory NH4+, overexpressing line S49-H and the isogenic control S49-NS were sprayed with the potent GS inhibitor, phosphinothricin (PPT; Gill and Eisenberg, 2001Go). To acquire an immediate and convenient measure of the physiological effect of PPT, we measured the photochemical efficiency of PSII, Fv/Fm. S49-NS plants sprayed with 0.2 mM PPT had a 27-fold increase in leaf NH4+ concentration compared with plants sprayed with water only (Fig. 5 ). An increase in NH4+ concentration was associated with a decrease in Fv/Fm, from an initial value of approximately 0.8, to below 0.6 at 0.2 mM PPT. The large pool of GDH isoenzyme 1 present in the leaves of line S49-H did not result in increased PPT resistance; there was almost identical NH4+ accumulation compared with S49-NS and a sharp decrease in photochemical efficiency (Fig. 5).


Figure 5
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  		Figure 5. Relative tolerance of a GDH beta-subunit overexpresser to the GS inhibitor, PPT. Plants were sprayed with PPT for 2 d. On the 3rd d, NH4+ concentration (white symbols, dashed lines) and the photochemical efficiency of PSII, Fv/Fm (black symbols, solid lines), were determined in the fourth true leaf of the control line, S49-NS (circles), and the GDH beta-subunit overexpressing line, S49-H (triangles). Data are the means ± SE (n = 6 for NH4+ concentrations and n = 14–18 for Fv/Fm). Error bars are not shown where smaller than the symbol and to improve clarity are sometimes shown in one direction only.

 

   DISCUSSION
TOP
 ABSTRACT
 RESULTS
 DISCUSSION
CONCLUSION
 MATERIALS AND METHODS
 LITERATURE CITED
 
Two different approaches were taken to study the in vivo reaction direction of GDH isoenzyme 1. First, we determined whether the increased amount of isoenzyme 1 present in the transgenic lines would result in an increase in [15N]Glu catabolism. Second, and conversely, we determined whether increased levels of isoenzyme 1 would result in an increased capacity to reassimilate photorespiratory NH4+ when the enzyme that usually has this role, GS, was inhibited.


GDH Isoenzyme 1 in Roots Catabolizes Glu

When [15N]Glu was supplied to roots of the control line, A63-NS, it was metabolized by at least three enzymes: GS, GAD, and GABA transaminase, producing labeled Gln amide, GABA, and Ala, respectively. Conceivably, the further metabolism of these labeled compounds could have produced the large amount of [15N]NH4+ that was also detected. However, when these enzymes were inhibited by the individual or combined action of MSX and AOA, a large amount of [15N]NH4+ still accumulated. Deaminative GDH activity would be a more straightforward origin for this [15N]NH4+, and therefore these results provide indirect evidence that the large amount of [15N]NH4+ produced by [15N]Glu-supplied roots resulted from substantial deaminative GDH activity.

Direct evidence for deaminative GDH activity in vivo is provided by the increased [15N]NH4+ accumulation rates observed in the GDH overexpressing lines. Compared to the control line, significantly more [15N]NH4+ accumulated in the overexpressing lines in both the control (65% increase over control line at 4 h) and MSX (27% increase) treatments (both P < 0.003), indicating that GDH isoenzyme 1 catabolizes Glu in roots. These results agree with those of Stewart et al. (1995)Go who supplied [15N]Glu via the roots to the maize GDH beta-subunit mutant and reported a 22-fold lower [15N]NH4+ concentration compared to a wild-type control in the presence of MSX. However, this study provides a more direct line of evidence, because the maize mutant had a 75% slower [15N]Glu uptake rate than a nonisogenic control, whereas in this study, the overexpressing lines had similar [15N]Glu uptake rates to an isogenic control.

The large intersample variation and consequent lack of significant differences among the genotypes (P > 0.05) observed in leaves is probably due to the relatively low levels of labeled metabolites in this tissue. Tobacco leaves have very low levels of GDH transcript, protein, and activity compared with roots (Purnell et al., 2005Go), and hence, the very low amount of [15N]NH4+ detected in leaves of the control line A63-NS was expected. Based on the results observed in roots, we expected that the large pool of GDH isoenzyme 1 present in leaves of the overexpressing lines would result in strong [15N]NH4+ accumulation. However, the overexpressing and control lines contained similar amounts of [15N]NH4+ in both the control and MSX treatments. A possible explanation for this result is that [15N]Glu supply to the leaves may have been small due to a slow nighttime transpiration rate, and, if so, this small amount of 15N[Glu] would have been diluted in a much larger pool of leaf [14N]Glu. Because GAD activity increases with increasing cytosolic Glu concentrations (Scott-Taggart et al., 1999Go), GAD-produced [15N]GABA was used as a marker of cytosolic [15N]Glu content. Considering that GAD activity is similar in tobacco roots and leaves (Baum et al., 1996Go), the fact that [15N]GABA accumulation in A63-NS leaves was only 7% of that in roots during the control treatment suggests that leaf [15N]Glu concentration was relatively low.


A Large Pool of Leaf GDH Isoenzyme 1 Is Incapable of Reassimilating Photorespiratory NH4+

In C3 plants such as tobacco, there is a large daytime flux of photorespiratory NH4+ within mesophyll cells (Keys et al., 1978Go). GS2 is crucial for the reassimilation of this NH4+, because mutants lacking this enzyme accumulate NH4+ and die (Wallsgrove et al., 1987Go). Although GDH activity is usually very low in mesophyll cells compared with GS2 (Bechtold et al., 1998Go), several studies have provided evidence that mesophyll GDH may reassimilate small amounts of photorespiratory NH4+ (Hartmann and Ehmke, 1980Go; Neeman et al., 1985Go; Yamaya et al., 1986Go). Moreover, transgenic tobacco plants expressing an NADP(H)-dependent GDH (EC 1.4.1.4) from Escherichia coli are partially resistant to GS-inhibiting PPT, accumulate more amino acids and approximately 10% more dry weight than wild-type plants when grown under NH4+ nutrition, produce 15N[Glu] when supplied 15N[NH4+] in the presence of PPT, and when supplied 13N[NH4+] produce more labeled Glu and Gln than controls (Ameziane et al., 2000Go; Mungur et al., 2005Go). Combined, these results provide strong evidence that the pool of E. coli GDH in these transformants reassimilates photorespiratory NH4+.

In the leaves of line S49-H, there is 34-fold more extractable NAD(H)-dependent GDH activity (EC 1.4.1.2) than in the leaves of the control line S49-NS (Purnell et al., 2005Go). Inhibition of GS2 activity by PPT should result in strong NH4+ accumulation in leaves. Therefore, if GDH isoenzyme 1 aminates 2-OG, the GDH overexpressing line S49-H should be partially resistant to PPT compared with the S49-NS control. Following treatment with PPT, line S49-H was no more resistant than line S49-NS, with the maximum effect of PPT at 0.20 mM. This contrasts with the transgenic plants overexpressing E. coli GDH, which were resistant to at least 7 times as much PPT (namely, >1.34 mM; Ameziane et al., 2000Go). Another notable difference is that the plants studied by Ameziane et al. (2000)Go grew larger under NH4+ nutrition compared with controls, whereas growth of S49-H and S4-H was not noticeably different from that of the control line A63-NS in hydroponic culture containing 1 mM NH4+ (Purnell et al., 2005Go). Because PPT does not inhibit GDH activity (Ameziane et al., 2000Go), these results suggest that, in contrast to E. coli GDH, plant GDH isoenzyme 1 does not assimilate NH4+ in vivo.


   CONCLUSION
TOP
 ABSTRACT
 RESULTS
 DISCUSSION
 CONCLUSION
MATERIALS AND METHODS
 LITERATURE CITED
 
Our results provide strong evidence that GDH isoenzyme 1 catabolizes Glu in roots, and does not assimilate NH4+ in source leaves. Fontaine et al. (2006)Go recently generated transgenic tobacco lines with decreased levels of the {alpha}-subunit and identified an Arabidopsis mutant similarly affected. Further work with these lines, as well as lines with perturbed levels of the beta-subunit (Purnell et al., 2005Go; Fontaine et al., 2006Go), is required to determine the in vivo reaction directions of isoenzymes 7 and 1, respectively, in different tissues at different developmental stages and in response to various environmental signals.


   MATERIALS AND METHODS
TOP
 ABSTRACT
 RESULTS
 DISCUSSION
 CONCLUSION
 MATERIALS AND METHODS
LITERATURE CITED
 

Plant Material and Growth Conditions

Tobacco (Nicotiana tabacum) L. var. Ti68 (McDaniel et al., 1996Go) seeds were germinated on a sandy loam in a growth chamber (16-h photoperiod, 150–300 µmol photons m–2 s–1, day/night temperature 25°C/18°C). Transgenic tobacco lines with modulated GDH activities used in this study were S4-H and S49-H (Purnell et al., 2005Go), which have 2- and 3-fold activity increases in the roots and 28- and 34-fold increases in the leaves, respectively, due to overexpression of a tomato (Solanum lycopersicum) GDH beta-subunit gene (Purnell et al., 1997Go). In addition, the NS lines A63-NS and S49-NS were used as isogenic controls.

For the [15N]Glu-labeling experiment, seedlings were transferred at 9 d postemergence (dpe) to an aerated hydroponic medium, as described previously (Purnell et al., 2005Go), except that the N source was 1 mM NH4NO3. At 20 dpe and 2 h into the dark period, the nutrient solution was replaced with an identical solution minus N and ± 1 mM MSX (Sigma) and/or ± 2 mM AOA (Sigma). The nutrient tanks were rocked gently to ensure adequate mixing. After 2 h, the solutions were replaced with identical solutions plus 5 mM [15N]Glu (95%–99% atom excess; Novachem). For the control and MSX treatments, six plants of each genotype were harvested at 2 and 4 h after the label was added (i.e. at the 6th and 8th h of the dark period). For the AOA and MSX + AOA treatments, six plants of S49-H and A63-NS were harvested at 4 h following addition of the label. At harvest, the roots of the plants were placed in 10 mM KCl/0.5 mM CaSO4 2H2O for 5 min to remove bound label and then rinsed twice in water. For each genotype, the plants were batched into three replicates of two plants per replicate. The roots and leaves of each pair of plants were diced finely, and root and leaf samples (approximately 0.5 g) were incubated overnight in 10 volumes ice-cold 100% methanol, before being stored at –20°C until needed.

For the NH4+ reassimilation test, seedlings were, at 15 and 16 dpe, sprayed to run off at the beginning of the photoperiod with a water-diluted PPT-containing herbicide (Basta, Hoechst; containing approximately 20% [v/v] PPT). Seventeen days postemergence and 30 min after the beginning of the photoperiod, the fourth true leaves of the seedlings were detached and dark adapted for 10 min. The photochemical efficiency of PSII, Fv/Fm, was then calculated with a Plant Efficiency Analyzer (Hansatech) operating at 60% maximum excitation light (1,400 µmol m–2 s–1) for 2 s. Subsequently, the leaves were incubated in 10 volumes ice-cold 100% methanol and the NH4+ concentration of the extracts determined by a colorimetric assay (McCullough, 1967Go). Because this assay is subject to interference from other metabolites (Schjoerring et al., 2002Go), the results presented here reflect relative differences between treatments but do not accurately report absolute NH4+ concentrations.


NMR Spectroscopy

Extracts from the [15N]Glu labeling experiment were reduced to dryness under vacuum and the pellets resuspended in 450 µL of a 0.2-M HCl/KOH buffer (pH 2.0; Weast, 1974Go). The solutions were clarified by centrifugation (13,000g, 4°C, 10 min) and then placed in 5-mm NMR tubes (Wilmad) with 50 µL of D2O (Aldrich).

NMR spectra were recorded on a Bruker Avance DMX750 spectrometer operating at 750 and 76 MHz for 1H and 15N, respectively, at 0°C, with a 5-mm 1H-13C-15N triple resonance probe equipped with triple axis gradients. All chemical shifts ({delta}) were referenced to TSP-d4 (Aldrich) and NH4Cl signals at {delta} = 0 ppm for 1H and 15N, respectively. Resonances were assigned by comparison with literature references (Fox et al., 1992Go) and with spectra of authentic 15N-labeled reference samples measured under identical experimental conditions to those employed for the sample extracts.

Routine 1H NMR spectra were acquired with water signal suppression using the WATERGATE pulse sequence (Sklenar et al., 1993Go) with an acquisition time of 1.99 s and a relaxation delay of 2.99 s. Free induction decays (FIDs) were acquired into 16 K complex data points with a spectral width of 8.25 kHz. Composite-pulse broadband 15N-decoupling (GARP) was used during the acquisition time for 15N-decoupled 1H spectra of labeled amino acid standards.

15N spectra were acquired using polarization transfer with a refocused INEPT pulse sequence (Burum and Ernst, 1980Go) using 16 K complex data points with a spectral width of 10.7 kHz and a repetition time of 3.5 s. The value of the 1JN-H evolutionary delay was set depending upon the targeted metabolite (74 and 91 Hz for NH4+ and Glu/Gln, respectively), and the refocusing delay was set to 1/81JN-H. 1H broadband decoupling (WALTZ16) was applied only during acquisition of the FID. Typically, 512 transients were acquired with a total data acquisition time of 30 min.

1H-15N reverse-correlation spectra were acquired using a one-dimensional version of the gradient-enhanced, sensitivity-improved HSQC pulse sequence (Kay et al., 1992Go). The 15N irradiation frequency was set to on resonance, and the 1JN-H evolutionary delays were optimized as above for each metabolite in separate experiments. The optimum refocusing delay was found to be 1/81JN-H and composite pulse broadband 15N decoupling (GARP) was applied during acquisition. A 1H trim pulse of 2 ms and gradient stabilization delays of 200 µs were employed. A total of 16 K complex data points were acquired using a spectral width of 8.25 kHz and repetition time of 2.5 s, requiring a total experiment time of 65 s/spectrum.

FIDs were Fourier transformed following exponential multiplication with a line-broadening factor of 0.5 Hz for 1H and 2 Hz for 15N spectra, with zero filling to 32 K complex data points. Peak areas were determined by line integration using the XwinNMR software program (Bruker).

For acquisition of 2D HSQC-TOCSY spectra (Bax and Davis, 1985Go; Davis et al., 1992Go), a 1/41JN-H evolution delay of 3 ms and a MLEV7 spin-lock mixing pulse of 80 ms with {gamma}·B1 = 25 kHz were employed. Composite pulse broadband 15N decoupling (GARP) was applied during acquisition. A total of 1,024 complex data points with 88 scans for each of 260 increments were acquired with spectral widths of 8.25 kHz and 7.6 kHz in F2 and F1, respectively. A repetition time of 2.12 s was used giving a total experiment time of 14.5 h. A {pi}/2-shifted sine-bell squared window function was applied with zero filling in each dimension to give a 2,048 x 256 complex data matrix following 2D Fourier transformation.


   ACKNOWLEDGMENTS
 
We thank Ian Brereton for expert assistance with NMR spectroscopy, David Anderson, Tony Cavallaro, and Yuri Trusov for technical assistance, and Scott Hermann for critical reading of the manuscript.

Received October 15, 2006; accepted November 3, 2006; published November 17, 2006.


   FOOTNOTES
 
1 Present address: Australian Institute for Bioengineering and Nanotechnology, The University of Queensland, c/o BSES Limited, 50 Meiers Road, Indooroopilly, QLD 4068, Australia. Back

The author responsible for distribution of materials integral to the findings presented in this article in accordance with the policy described in the Instructions for Authors (www.plantphysiol.org) is: José Ramon Botella (j.botella{at}uq.edu.au).

[OA] Open Access articles can be viewed online without a subscription. Back

www.plantphysiol.org/cgi/doi/10.1104/pp.106.091330

* Corresponding author; e-mail j.botella{at}uq.edu.au; fax 61–7–3365–1699.


   LITERATURE CITED
TOP
 ABSTRACT
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 DISCUSSION
 CONCLUSION
 MATERIALS AND METHODS
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