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Plant Physiol. Nov 2002; 130(3): 1263–1275.
PMCID: PMC166647

Functional Analysis of an Arabidopsis T-DNA “Knockout” of the High-Affinity NH4+ Transporter AtAMT1;11

Abstract

NH4+ acquisition by plant roots is thought to involve members of the NH4+ transporter family (AMT) found in plants, yeast, bacteria, and mammals. In Arabidopsis, there are six AMT genes of which AtAMT1;1 demonstrates the highest affinity for NH4+. Ammonium influx into roots and AtAMT1;1 mRNA expression levels are highly correlated diurnally and when plant nitrogen (N) status is varied. To further investigate the involvement of AtAMT1;1 in high-affinity NH4+ influx, we identified a homozygous T-DNA mutant with disrupted AtAMT1;1 activity. Contrary to expectation, high-affinity 13NH4+ influx in the amt1;1:T-DNA mutant was similar to the wild type when grown with adequate N. Removal of N to increase AtAMT1;1 expression decreased high-affinity 13NH4+ influx in the mutant by 30% compared with wild-type plants, whereas low-affinity 13NH4+ influx (250 μm–10 mm NH4+) exceeded that of wild-type plants. In these N-deprived plants, mRNA copy numbers of root AtAMT1;3 and AtAMT2;1 mRNA were significantly more increased in the mutant than in wild-type plants. Under most growth conditions, amt1;1:T-DNA plants were indistinguishable from the wild type, however, leaf morphology was altered. However, when grown with NH4+ and sucrose, the mutant grew poorly and died. Our results are the first in planta evidence that AtAMT1;1 is a root NH4+ transporter and that redundancies within the AMT family may allow compensation for the loss of AtAMT1;1.

In many natural ecosystems and in high-input monoculture cropping systems, nitrogen availability represents a limiting factor, necessitating the use of large quantities of nitrogen fertilizers (presently approximately 1011 kg per annum globally; http://www.fao.org/) to achieve current high yields of quality crops (Marschner, 1995). However, this high level of nitrogen fertilizer use may generate environmental problems, such as ground water contamination by nitrate, anoxia of rivers and oceanic coastal waters, or even fish kills attributable to leaching of N from agricultural soils (Cohen et al., 1996). In cereal crops, which receive roughly 60% of all N fertilizers, recovery of applied N in grain is typically only 33% of fertilizer applied to soils (Raun and Johnson, 1999). This low N use efficiency is caused by several factors, including volatilization of NH3, denitrification, and/or leaching of soil nitrate (NO3). However, it is also the result of inefficient N absorption (KM values for NO3 and NH4+ absorption by higher plants may be orders of magnitude higher than those of algae and fungi (Galvan et al., 1996; Unkles et al., 2001). Furthermore, as plants accumulate N, there is a rapid down-regulation of genes encoding high-affinity NO3 and NH4+ transporters (Rawat et al., 1999; Forde, 2000; Howitt and Udvardi, 2000; Glass et al., 2001), and there are corresponding reductions of high-affinity influx (Glass and Siddiqi, 1995; Gazzarrini et al., 1999; Rawat et al., 1999; Zhuo et al., 1999; von Wiren et al., 2000a). In addition, as external N levels increase, N efflux from roots becomes a significant proportion of net influx, especially in the case of NH4+, where futile NH4+ cycling (leak and pump across the plasma membrane) has been demonstrated to increase respiration rates by 40% in barley (Hordeum vulgare) roots (Britto et al., 2001). It clearly will be essential to better define the mechanisms involved in regulating N acquisition, assimilation, and redistribution in the plant to enable agronomists to improve the management of plant N nutrition and to achieve genetic modifications that might sustain present high rates of crop productivity with reduced N inputs.

A first step in improving plant N use efficiency would be to address the primary acquisition process occurring at the root/soil interface. Genes encoding NO3 transporters, NRT1 and NRT2 (Tsay et al., 1993; Forde, 2000) and NH4+ transporters, AMT1 and AMT2 (Ninnemann et al., 1994; Lauter et al., 1996; Gazzarrini et al., 1999; Sohlenkamp et al., 2000; von Wiren et al., 2000a) have been identified in plants. These transporters are presumed to be located mainly at the root plasma membrane and, therefore, positioned to take N up from the soil. Nevertheless, in the cases of both NO3 and NH4+ uptake, many more genes have been identified in Arabidopsis and other organisms than would have been anticipated from physiological studies (Glass and Siddiqi, 1995; Crawford and Glass, 1998; Forde and Clarkson, 1999). An important challenge is to ascribe specific functions to each of these genes.

Although NO3 is the predominant form of N in well-aerated and pH-balanced soils, NH4+ is an important and commonly underestimated N source for many plant species, in part because in mixtures of NO3 and NH4+, NH4+ inhibits NO3 uptake. In Lemna sp., in rice (Oryza sativa), and in Arabidopsis, NH4+ influx consists of both saturable high-affinity influx (HATS) and non-saturable low-affinity influx (LATS) that operate at low and high NH4+ concentrations, respectively (Ullrich et al., 1984; Wang. et al., 1993; Rawat et al., 1999). It is still unknown what molecular mechanisms are involved in either the high- or low-affinity NH4+ transport phenomenon in plant roots.

Some members of the AMT1 family are suggested to be putative high-affinity NH4+ transport proteins in planta based on their functional analysis when expressed in yeast cells (see below). Included in this group of genes are AtAMT1;1, AtAMT1;2, and AtAMT1;3 (Gazzarrini et al., 1999; Rawat et al., 1999) and three AMT1 homologs in tomato (Lycopersicon esculentum; Lauter et al., 1996; von Wiren et al., 2000a). Two other AMT1 homologs, AtAMT1;4 and AtAMT1;5, have been identified with the complete sequencing of the Arabidopsis genome; whether they are involved in NH4+ transport remains to be determined (von Wiren et al., 2000b). A tomato homolog, LeAMT1;1, found in root hairs has recently been expressed in Xenopus sp. oocytes and shown to function as an NH4+ uniporter dependent on membrane potential and NH4+ concentration gradients (Ludewig et al., 2002).

Sequencing of the Arabidopsis genome has also resulted in the identification of a second class of AMT genes designated AMT2 (Sohlenkamp et al., 2000). Functional expression of AtAMT2 in yeast cells revealed a strikingly different phenotype to that of AMT1, where the uptake of NH4+ resembled a low-capacity transport system (Sohlenkamp et al., 2000). Members of the AMT gene families show distinctive patterns of RNA expression in roots and shoots and during the diurnal cycle (Gazzarrini et al., 1999; Rawat et al., 1999; von Wiren et al., 2000a), which suggests that this gene family may serve a number of different functions associated with NH4+ transport across the plasma membrane and within the plant.

AtAMT1;1 has traditionally been considered of prime importance in Arabidopsis NH4+ transport, particularly in the roots where its mRNA is highly abundant. High-affinity NH4+ influx is strongly correlated with transcript abundance, which appears to be regulated by root Gln (but not NH4+) concentration (Gazzarrini et al., 1999; Rawat et al., 1999; Gansel et al., 2001). Furthermore, functional analysis in yeast cells demonstrated that AtAMT1;1 is capable of mediating high-affinity NH4+ transport with a significantly higher affinity for NH4+ (KM approximately 500 nm) than its counterparts AtAMT1;2 and AtAMT1;3 (KM approximately 40 μm; Gazzarrini et al., 1999; Shelden et al., 2001). To decipher the role of AtAMT1;1 in Arabidopsis, we have used a reverse genetic approach to identify an amt1;1 transfer-DNA (amt1;1:T-DNA) tagged line. Disruption of AtAMT1;1 activity reduced high-affinity NH4+ influx by only 30%, whereas low-affinity transport was increased. Additional phenotypic effects of this disruption included an altered leaf morphology and a lethal condition in plants grown under sterile conditions in the presence of NH4+ and Suc.

RESULTS

amt1;1 T-DNA Isolation and Characterization of Inserted Loci

The amt1;1:T-DNA line was identified from the subpool, CD6-4A, of the “Jack enhancer-trap” lines. DNA isolated from individual lines was screened using the PCR with the T-DNA right border primer (P3) and the AtAMT1;1 gene specific primer (P2; Table TableI).I). A PCR fragment of the expected size was identified in which the right border of the T-DNA was inserted 6 bp upstream of the first putative Met in the open reading frame of AtAMT1;1 (Fig. (Fig.1A).1A). Inverse PCR revealed that the left border was inserted 79 bp upstream of the right border insertion, deleting a 72 bp fragment of the genome (Fig. (Fig.1A).1A). A 3-bp insertion of unknown DNA was also inserted in this junction (Fig. (Fig.1A).1A). The amt1;1:T-DNA line was selfed and then backcrossed to Col3 gl1. Because no visible phenotype was apparent in the mutant (see below) homozygous lines were selected using both a PCR-based screen and Southern-blot analysis.

Table I
List of oligonucleotides used in this study
Figure 1
Characterization of the plant line with a T-DNA insertion in AtAMT1;1. A, Localization of the T-DNA insert. The diagram illustrates the insertion of the T-DNA 6 bp upstream of the first putative Met in the open reading frame of AtAMT1;1. The bordering ...

DNA isolated from the backcrossed lines were subjected to two rounds of the PCR. The first round was designed to identify the T-DNA-tagged amt1;1 allele using primers to amplify the 1.7-kb region spanning the right border (P3) of the T-DNA insert and the 3′ end of AtAMT1;1 (P7; Fig. Fig.1A).1A). The second PCR used primer pairs (P8 and P7) to identify the wild-type allele, amplifying a 2.3-kb region of genomic DNA, which included the upstream promoter region and the 3′ end of AtAMT1;1 (Fig. (Fig.1A).1A). A homozygous line that failed to amplify the wild-type allele but did amplify the T-DNA-tagged amt1;1 was selected and selfed (data not shown). To identify the number of T-DNA insertion events, Southern-blot analysis was performed on EcoRI or HindIII digested genomic DNA. Using a 1.2-kb cDNA probe specific to the uidA (GUS) open reading frame, single DNA fragments of 15 and 5.4 kb were identified in the EcoRI- and HindIII-digested DNA, respectively (Fig. (Fig.1C).1C). HindIII restriction sites are present close to the right border of the T-DNA and approximately 400 bp upstream from the left border T-DNA insertion site in chromosome 4 (Fig. (Fig.1B).1B). As expected, digestion with HindIII liberated a 5.4-kb DNA fragment from the AMT1;1 loci, which contained the majority of the T-DNA insert. Digestion with EcoRI also confirmed the presence of a single-insertion event in amt1;1:T-DNA (Fig. (Fig.1C),1C), however, the identified 15-kb fragment was smaller than the predicted fragment size of 20 kb based on a restriction enzyme digest profile of the bacterial artificial chromosome sequence (accession no. AL049656).

Disruption of AtAMT1;1 mRNA Expression

The expression of AtAMT1;1 was examined using northern-blot analysis in both amt1;1:T-DNA and wild-type plants maintained on 1 mm NH4NO3 for 5 weeks and then deprived of N for 4 d. This treatment up-regulates AtAMT1;1 mRNA expression in roots (Gazzarrini et al., 1999; Rawat et al., 1999; Gansel et al., 2001). Northern blots containing total RNA from both the T-DNA mutant and wild-type roots and shoots were incubated with a 1.7-kb DIG-labeled AtAMT1;1 anti-sense RNA probe. A 1.7-kb hybridization product was identified in the wild-type root and shoot total RNA extracts but not in the T-DNA mutant (Fig. (Fig.11D).

Growth Analysis

No discernible phenotype was evident when T-DNA mutants were grown on a peat-based soil or in an open hydroponic system in nutrient solution with no added C. Plants germinated, bolted, and set flower as did wild-type plants. When grown in open hydroponic systems for 6 weeks on high-N media containing 1 mm NH4NO3 or 2 mm KNO3, or low-N media containing 50 μm (NH4)2SO4 or 100 μm KNO3, both without added C, no significant differences in fresh weights were detected (Table (TableII).II). By stark contrast, mutant plants grown in sterile hydroponic conditions with 1% (w/v) Suc and 0.5 mm (NH4)2SO4 grew poorly relative to the wild type and died after 2 to 3 weeks (Fig. (Fig.2,2, A and B). This lethality was not expressed when 0.5 mm (NH4)2SO4 was replaced by 1 mm KNO3 under otherwise identical conditions and equivalent N levels (Fig. (Fig.2C).2C).

Table II
Growth analysis of wild-type and amt1;1:T-DNA plants
Figure 2
Plant growth in magenta boxes containing sterile nutrient solution including Suc. Stratified seeds were sown directly onto nylon mesh sitting on the floating rafts in nutrient solution containing 1% (w/v) Suc and either 0.5 mm (NH4)2SO4 (A and ...

Leaves of the amt1;1:T-DNA were found to be distinctly more succulent than those of the wild type. They were characterized by a thickening of the mesophyll tissue along the major vein of the leaf, and a significant decrease in the extent of intercellular air spaces between blade mesophyll cells (Fig. (Fig.3).3).

Figure 3
Cross sections of wild-type and amt1;1:T-DNA leaves. Sections are from leaves of plants grown in a controlled temperature growth chamber in hydroponic tanks (A) or in the greenhouse (B). Plants grown in the greenhouse were on a peat-based potting mix ...

The Root High-Affinity NH4+ Transport System

Rates of high-affinity 13NH4+ influx into intact roots of N-replete wild-type and amt1;1:T-DNA plants were similar (Fig. (Fig.4A).4A). However, after 4 d of N deprivation, significant differences in 13NH4+ influx and Vmax values for influx were apparent whereas trends toward higher KM values were evident in the amt1;1:T-DNA over those of the wild type. KM and Vmax values were 17.2 ± 4.4 μm and 8.4 ± 0.5 μmol 13NH4+ g−1 fresh weight h−1 for amt1;1:T-DNA and 11.3 ± 3.2 μm and 11.4 ± 0.6 μmol 13NH4+ g−1 fresh weight h−1 for the wild type (Fig. (Fig.4B).4B).

Figure 4
13NH4+ influx by the HATS for both amt1:1:T-DNA and wild-type plants. A, Plants were grown in the presence of 1 mm NH4NO3 for 5 weeks and then transferred to nutrient solution with or without 1 mm NH4NO3 for 4 d. 13NH4+ influx was measured ...

The Root Low-Affinity NH4+ Transport System

At higher NH4+ concentrations (up to 10 mm), 13NH4+ influx (the sum of LATS and HATS), was higher in the mutant than in the wild type (Fig. (Fig.5A).5A). This was even more apparent when the calculated HATS Vmax values were subtracted from the combined LATS and HATS activities to give the corrected LATS flux (Fig. (Fig.5B).5B).

Figure 5
NH4+ influx by the LATS for both amt1;1:T-DNA and wild-type plants. Both sets of plants were grown on 1 mm NH4NO3 for 5 weeks and then transferred to nutrient solution without N for 4 d. A, Concentration dependence of 13NH4+ influx by ...

Root and Shoot Tissue N Analysis

Analyses of root and shoot NH4+, NO3, Gln, and %N and %C revealed only minor differences (that were not statistically significant) between the amt1:1:T-DNA and wild-type lines regardless of N status (Table III). After 4 d of growth without exogenous N, %N decreased resulting in elevated C to N ratios in the shoots and roots of both lines.

Table III
Root and shoot nitrogen and carbon pools from wild-type and amt1;1:T-DNA plants grown hydroponically for 5 weeks in the presence of 1 mm NH4NO3 (+N) or after a 4-d period where nitrogen was removed from the nutrient solution (−N)

Competitive Reverse Transcriptase (RT)-PCR: Profile of mRNA Expression Patterns by the AtAMT1 and AtAMT2 Gene Families

Copy numbers of AtAMT mRNA transcripts in roots of wild-type and mutant plants maintained continuously in 1 mm NH4NO3 were obtained using competitive RT-PCR (for a typical reaction, see Fig. Fig.6A).6A). In wild-type plants, gene expression levels were in the order AtAMT1;2> AtAMT1;1> AtAMT1;3 > AtAMT2;1 (Fig. (Fig.6C).6C). This hierarchy was relatively unchanged after 4 d of N deprivation, but copy numbers increased by approximately 1.9-, 1.5-, 1.9-, and 1.3-fold for AtAMT1;2, AtAMT1;1, AtAMT1;3, and AtAMT2;1 respectively. These increases were statistically significant (P < 0.05) for AtAMT1;1, AtAMT1;2, and AtAMT1;3. In the amt1;1:T-DNA line, AtAMT1;2 was also the most abundantly expressed AMT followed in order by AtAMT1;3 and AtAMT2;1. N deprivation increased AtAMT1;2, AtAMT1;3, and AtAMT2;1 expression by 1.6, 2.6, and 1.6 times, respectively. Interestingly for AtAMT1;3 and AtAMT2;1, these increases were significantly greater (P < 0.05) than the corresponding increases in wild-type plants (Fig. (Fig.6C).6C).

Figure 6
Quantitative AMT gene expression levels estimated using competitive RT-PCR. Plants were grown on 1 mm NH4NO3 for 5 weeks and then transferred to nutrient solution with (T = 0) or without (T = 4) N for 4 d before harvest of root tissues. A, A typical multiplex ...

DISCUSSION

amt1;1:T-DNA was identified from a set of enhancer-trap T-DNA tagged Arabidopsis lines in the Col background (Campisi et al., 1999). Sequence analysis revealed that the right border of the T-DNA tag was inserted 6 bp upstream of the first putative ATG of the open reading frame of AtAMT1;1 (Fig. (Fig.1A).1A). Although the T-DNA tag is not inserted within the open reading frame of AtAMT1;1, its presence directly upstream of the coding sequence was sufficient to disrupt the transcription of AtAMT1;1 (Fig. (Fig.1D).1D). No AtAMT1;1 mRNA transcripts were found in roots or shoots of amt1;1:T-DNA plants deprived of N for 4 d, a treatment, which resulted in high levels of AtAMT1;1 expression in wild-type plants (Fig. (Fig.1D).1D). Failure to identify AtAMT1;1 mRNA supports the initial genome analysis using the PCR that our amt1;1:T-DNA is a homozygous line. However, the observed reduction in expression of AtAMT1;1 could also result from deleterious insertion or deletion by other T-DNA insertions elsewhere in the genome acting in trans to silence AtAMT1;1. Southern-blot analysis using a GUS DNA probe, which would identify other T-DNA insertions, revealed that there was only a single T-DNA insertion in the amt1;1:T-DNA genome (Fig. (Fig.11C).

The amt1;1:T-DNA and wild-type plants grown on adequate N (1 mm NH4NO3) showed similar rates of 13NH4+ flux into roots in the high-affinity range (25 and 100 μm; Fig. Fig.4A).4A). The introduction of a 4-d N starvation period increased 13NH4+ influx into roots for both lines (Fig. (Fig.4,4, A and B), however the flux in the amt1;1:T-DNA line was generally only 30% to 40% lower than in roots of wild-type plants. This phenotype was also present with plants grown on 2 mm KNO3 in the presence of 1% (w/v) Suc followed by a 4-d N starvation period (data not shown). Derepression of root NH4+ influx by N starvation is consistent with previous results in Arabidopsis (Gazzarrini et al., 1999; Rawat et al., 1999) and in rice (Wang et al., 1993). Failure to more substantially reduce NH4+ influx after disrupting AtAMT1;1 suggests that this gene may encode a transporter that normally contributes only 30% to 40% of high-affinity NH4+ influx, despite its suggested higher affinity for NH4+ than other AtAMT1 transporters (Gazzarrini et al., 1999); certainly mRNA levels of AtAMT1;2 were actually higher than those of AtAMT1;1. As an alternative, AtAMT1;1 may quantitatively be a major player in high-affinity NH4+ influx, and other NH4+ transporters partially compensate for its disruption in the mutant. Estimated KM values for 13NH4+ influx were 11.3 ± 3.2 μm and 17.2 ± 4.4 μm for the wild type and amt1;1:T-DNA, respectively. Although not significantly different, there is a trend toward a higher KM in the amt1;1:T-DNA line than the wild type, which is consistent with the estimated lower KM values for AtAMT1;1 in yeast (approximately 0.5 μm), compared with values of approximately 40 μm for AtAMT1;2 and AtAMT1;3 (Gazzarrini et al., 1999). However, it is possible that the loss of AtAMT1;1 and its contribution to the plant's combined KM for NH4+ is being masked by the apparent compensation of other AMT proteins and possibly by other yet unidentified NH4+ transporters. Our estimates of KM values are substantially lower than were reported in another study in Arabidopsis (Rawat et al., 1999), which also used 13NH4+ to measure influxes as a function of external NH4+ concentration. These differences may be attributable to the use of sterile growth conditions within Magenta boxes or to the presence of Suc in the growth media in the study by Rawat et al. (1999). However, it was noted by Wang et al. (1993) that KM values for NH4+ influx in rice roots varied with N status of the plants.

When 13NH4+ influx was measured at higher concentrations of NH4+ (1–10 mm) in plants that had been deprived of N for 4 d, influx of 13NH4+ through the low-affinity system had increased in the amt1;1:T-DNA line relative to wild-type plants (Fig. (Fig.5).5). This increased flux by the low-affinity system may be the result of plant compensation for the reduction of high-affinity NH4+ flux caused by the loss of AtAMT1;1. Our earlier studies and those of other groups (see Glass and Siddiqi, 1995) have noted the negative correlations between high-affinity NH4+ influx and accumulated N. In the absence of a functional AtAMT1;1 transporter, the reduced tissue N may result in elevated expression of mRNAs encoding all transporters that are regulated transcriptionally by tissue N and in particular Gln (Rawat et al., 1999). This may account for the increased LATS flux of 13NH4+ in the present study and the maintenance of NH4+ pools and %N in the mutant (Table III). However, the well-documented reduction of high-affinity NH4+ influx associated with high-N tissue status failed to apply to low-affinity transport in rice, where 13NH4+ influx attributable to LATS was increased in plants previously grown on 1 mm NH4+ compared with plants grown on 2 μm or 100 μm NH4+ (Wang et al., 1993). The apparent failure to down-regulate the LATS after growth on NH4+ was also observed in roots of aspen (Populus spp.), lodgepole pine (Min et al., 2000), and citrus (Cerezo et al., 2001), even though the HATS was significantly down-regulated by this treatment. This anomaly is counterintuitive given that the HATS was down-regulated under these conditions and remains unexplained, but our present observation with the mutant may represent a similar phenomenon, whereby suppression of HATS is associated with increased influx through the LATS. Britto et al. (2001) suggest that at elevated NH4+, which would repress the activity of the HATS, there is a failure to regulate LATS influx, resulting in an energetically costly active efflux of NH4+ and a massive futile cycling across the plasma membrane. This may in part explain the toxic effects of elevated external NH4+ on many plant species (Van der Eerden, 1982; Kronzucker et al., 2001). This observed increase of LATS flux suggests some form of interaction between the high- and low-affinity NH4+ uptake systems, possibly through enhanced expression of NH4+ regulated high-affinity AMT genes including AtAMT1;2, AtAMT1;3, and AtAMT2;1 (Gazzarrini et al., 1999; Shelden et al., 2001; this study). Shelden et al. (2001) have reported that AtAMT1;2 expressed in yeast displays biphasic kinetics, accumulating 14C-methylammonium at both low (0–100 μm) and high (0.25–10 mm) concentrations. It is possible that the low-affinity NH4+ flux observed in this study may be the result of a switch in protein activity and NH4+ affinity by AtAMT1;2 (Shelden et al., 2001). However, with the loss of AtAMT1;1 activity, AtAMT1;2 gene expression did not increase relative to the wild type as did AtAMT1;3 and AtAMT2;1, and therefore, from a mRNA transcript perspective, it appears less likely that AtAMT1;2 is the primary contributor to the increase in LATS flux in the amt1;1:T-DNA line. However as mentioned above, we cannot rule out a switch in AtAMT1;2 transport activity that may have occurred in the mutant in response to the loss of AtAMT1;1 activity. The protein AtAMT2;1, which is distantly related to members of the AMT1 family in Arabidopsis, has recently been identified and characterized as a putative low-capacity NH4+ transporter (Sohlenkamp et al., 2000). AtAMT2;1 is expressed in both shoots and roots and responds to periods of N starvation (Sohlenkamp et al., 2000). Whether its activity is related to the increased flux of NH4+ in the low-affinity range in the amt1;1:T-DNA line remains to be shown. However, as discussed below, its expression is enhanced in the amt1;1:T-DNA relative to the wild type making it a good candidate for part of this NH4+ transport phenomenon.

To investigate the hypothesis that disruption of AtAMT1;1 was being compensated for by overexpression of other members of the AMT family we made use of competitive RT-PCR to measure total mRNA copy numbers and directly compare expression patterns among members of the AtAMT1 and AtAMT2 families. As expected, wild-type plants subjected to a 4-d period of N starvation increased levels of expression of all genes examined relative to plants grown with sufficient N over that period. This is a result that is consistent with other studies that have demonstrated the strong response of some members of the AMT family (AtAMT1;1 and AtAMT1;3) to N starvation (Gazzarrini et al., 1999; Rawat et al., 1999; Sohlenkamp et al., 2000; Gansel et al., 2001; Shelden et al., 2001). Interestingly, using the sensitive competitive RT-PCR technique we were able to observe consistent increases in AtAMT1;2 mRNA levels that have not been previously observed by other experimenters using total RNA northern blots (Gazzarrini et al., 1999; Shelden et al., 2001).

In wild-type plants, the order in which the AMT genes were expressed in roots after N starvation from most to least abundant was AtAMT1;2 > AtAMT1;1 > AtAMT1;3 > AtAMT2;1 (Fig. (Fig.6C).6C). Apart from AtAMT1.1, this same pattern was evident in the amt1;1:T DNA line, however, levels of AtAMT1;3 and AtAMT2;1 were significantly higher than were observed in the wild type. We suggest that this elevated level of gene expression may have increased NH4+ transport capacity through other AMT transporters, which has the effect of masking the loss of the contribution of AtAMT1;1 and which explains the small reduction in NH4+ transport observed in the amt1;1:T-DNA line. This raises an important consideration vis à vis the effectiveness of the strategy of using particular gene mutations to determine the role of those genes. Where there is the capacity to compensate for the loss of activity, the role (particularly at the quantitative level) of disrupted genes may be obscured. The apparent compensation by AtAMT1;3 and AtAMT2;1 also raises the question of the mechanism of this putative compensation. The simplest explanation would be a form of passive compensation. Because the complete absence of AtAMT1;1 transporters should significantly reduce N uptake and tissue N concentration, down-regulation of all genes regulated by tissue N levels should be reduced, and these genes will consequently be overexpressed relative to their expression levels in wild-type plants.

By contrast, in a study of high-affinity NO3 transport, there was a lack of compensation for disruption of the two high-affinity NO3 transport systems AtNRT2;1 and AtNRT2;2 (Filleur et al., 2001). A T-DNA tagged mutant disrupted in two genes encoding the high-affinity NO3 transporters AtNRT2;1 and AtNRT2;2 were shown to have a Vmax for 15NO3 influx that was only 27% of wild-type rates, with no apparent changes of flux values in the low-affinity range (Filleur et al., 2001). Because two genes have been inactivated in this line, their individual contributions to NO3 flux may have exceeded any compensation in NO3 uptake by other NRT2 proteins. As an alternative, it is possible that only AtNRT2;1 and AtNRT2;2 encode transporters involved in high-affinity NO3 influx into roots, the other AtNRT2 genes possibly participating in internal redistribution of absorbed NO3 such as NO3 fluxes to the stele. It is also possible that NO3 regulatory elements may have been disrupted that could limit subsequent activation of the NRT2 system during the synthesis of the T-DNA mutant, because 25 kb of genomic DNA was deleted with the T-DNA insertion. However, in another study of NO3 transport in Aspergillus nidulans where there appear to be only two members of the NRT family and no other NO3 transporters, loss of the high-capacity NRTA (formerly CRNA) gene reduced NO3 influx to approximately 10% of wild-type levels with no apparent compensation by the second gene NRTB (Unkles et al., 2001).

There was no readily apparent phenotype in the amt1;1:T-DNA line when grown in soil in growth chambers under light conditions of approximately 150 to 200 μmol m−2 s−1 (B.N. Kaiser, unpublished data). Using an open-top hydroponic system, plants were grown on low (100 μm NH4+ or KNO3) or adequate (1 mm NH4NO3 or 2 mm KNO3) N without external C under the same conditions used in the 13NH4+ influx studies. Regardless of N form (NH4+ or NO3) or concentration, after 5 weeks there were little differences in fresh weights of roots and shoots between the amt1;1:T-DNA and the wild type with the exception that amt1;1:T-DNA plants grown in 100 μm KNO3 had lower root and shoot fresh weights than wild-type plants (Table (TableII).II). Correspondingly, %N, NO3, NH4+, and Gln levels in roots and shoots of the wild-type and amt1;1:T-DNA lines were similar (Table III).

Although little differences were observed in fresh weights, leaves of the T-DNA mutant were more succulent to the touch. Analysis of embedded cross sections of various aged leaves identified a developmental response with age (B.N. Kaiser, unpublished data) of increased cell density and reduction of intercellular air spaces specifically around the central vein, which was enlarged (Fig. (Fig.3).3). This phenotype was present in both hydroponically grown plants in a growth-chamber and in soil grown greenhouse plants both supplied nutrient solution with adequate levels of N (1 mm NH4NO3). Although we saw little difference between the wild-type and the amt1;1:T-DNA lines in 13NH4+ influx into roots of plants supplied N, transport properties within the shoots could be entirely different. The loss of AtAMT1;1 activity may influence translocation of NH4+ within leaves and the loading or unloading of NH4+ into the vascular cylinders. The balance between NH4+ and NO3 in leaves has been suggested to influence leaf growth and cell division through indirect effects on cellular cytokinin levels and osmoticum balance (Walch-Liu et al., 2000). Although NO3 pool sizes were similar in the shoot and roots of the amt1;1:T-DNA and wild-type lines, there were differences in the balance of NO3 to NH4+ pools (Table (TableII).II). The marginally higher ratio of NO3 to NH4+ in the shoots of the amt1;1:T-DNA line may suggest that cells adjacent to the central vein are experiencing altered N ratios (NO3>NH4+) and possibly influencing localized cell development. Subsequent studies are planned to investigate the role of AtAMT1;1 on NH4+ transport within leaves and its possible involvement in the regulation of cell division and N remobilization between developing plant organs.

Arabidopsis is often grown under sterile conditions in the presence of Suc (Gazzarrini et al., 1999; Rawat et al., 1999). In the presence of 1% (w/v) Suc and 0.5 mm (NH4)2SO4, the amt1;1:T-DNA would germinate and produce their first set of leaves; but then during the 10 d after germination, plants grew poorly with little new leaf or root production (Fig. (Fig.2,2, A and B). By contrast, when 0.5 mm (NH4)2SO4 was replaced by 1 mm KNO3, amt1;1:T-DNA grew essentially as the wild-type plants (Fig. (Fig.2C).2C). Growth of the amt1;1:T-DNA line with NH4+ and Suc also resulted in reddening of the leaves caused by the accumulation of anthocyanins (Fig. (Fig.2A),2A), an Arabidopsis phenotype that has been associated with stress responses including nutrient deficiencies and excess carbohydrate (Dangl, 1991; Martin et al., 2002). Martin et al. (2002) have recently demonstrated that the C to N ratio in Arabidopsis seedlings may regulate growth through possible effects on the mobilization of seed storage reserves and photosynthetic gene expression. Poor root and cotyledon development and accumulation of anthocyanins was significant in Arabidopsis seedlings grown in the presence of high levels of Suc (100 mm) and low N concentrations (100 μm; Martin et al., 2002).

It is interesting to speculate on the negative effects of NH4+ provision in the presence of exogenous Suc in the amt1;1:T-DNA line. AtAMT1;1 mRNA expression has been examined previously under conditions where Suc was present in (Rawat et al., 1999) or absent from (Gazzarrini et al., 1999) the growth media. In the presence of Suc, AtAMT1;1 expression increased (up to 10-fold) and decreased dramatically within very short time periods (<24 h) in response to N starvation and resupply (Rawat et al., 1999). By contrast without Suc, the rate of change in AtAMT1;1 expression was much less rapid. In this study using competitive RT-PCR, we observed a 1.5-fold stimulation of AtAMT1;1 expression after a 4-d starvation period, whereas using traditional northern-blot analysis, Gansel et al. (2001) reported a 2-fold stimulation after a 5-d N starvation period, and Gazzarrini et al. (1999) a 3-fold increase after a 3-d N starvation period. The data of Rawat et al. (1999) suggests that Suc attenuates the activation of AtAMT1;1 expression. The mechanisms involved are unknown, however, it is possible that Suc may be acting as a component of a signaling cascade involved in maintaining C to N balance in plant tissues, with AtAMT1;1 possibly serving as the primary receptor involved in N sensing and NH4+ transport per se. Although, the influx data demonstrated that 13NH4+ influx through the LATS system was actually increased in the amt1;1:T-DNA line, and there were also compensatory effects on the expression of other members of the AMT family, the reduced influx of NH4+ at moderate external concentrations (0–1 mm), which was also apparent in Suc grown plants (data not shown), may have been sufficient to disturb the C to N balance in the young developing seedlings resulting in the observed toxic effects of poor seedling development, root growth, and accumulation of anthocyanins.

In summary, we have generated a T-DNA mutant disrupted in the expression of AtAMT1.1. This mutation resulted in a reduction of approximately 30% to 40% in high-affinity root NH4+ transport under N-limiting conditions. Albeit the reduction in high-affinity NH4+ transport is less than initially expected from the strong correlative results of AtAMT1;1 expression and NH4+ transport published by this group (Rawat et al., 1999) and others (Gazzarrini et al., 1999; Gansel et al., 2001), this is the first documented evidence, to our knowledge, that a member of the AMT family is an NH4+ transporter in planta involved in NH4+ uptake into plant roots. The failure to observe a more pronounced reduction in high-affinity NH4+ transport appears to be the result of compensation by overexpression of other members (AtAMT1;3 and AtAMT2;1) of the AMT families of genes. Internal N pools and %N in roots and shoots were similar for the amt1;1:T-DNA and the wild-type lines. The results demonstrate that genetic redundancy in plants, in the case of the AMT family, provides an important ecological plasticity capable of adapting to mutation in nature. Subsequent analysis of other individual and multiple AMT disrupted mutants will help to resolve the question of the functions of each AMT gene and help to explain the compensatory phenomenon observed in this study. There are, nevertheless, as yet unexplained interactions leading to altered leaf morphology in the mutant and the significant interaction between C supply (in the form of Suc) and NH4+ transport in the mutant, resulting in a highly lethal phenotype. These two phenotypes provide an exciting opportunity to examine the dynamic control of cell development by nutritional status controlled by the AMT family of NH4+ transport proteins.

MATERIALS AND METHODS

Plant Growth

Arabidopsis Col-3 gl1 (wild type) and amt1-1:T-DNA (mutant) were grown either in a controlled chamber with a 25°C/20°C day/night temperature regimen in a hydroponic system (see below) or in peat-based soil media. Plants were illuminated by Vita-lite fluorescent lamps (Durotest, Fairfield NJ), which generated between 150 and 200 μmol m−2 s−1 of photosynthetically active radiation at plant level. During T-DNA isolation and subsequent seed multiplication and background crosses, plants were grown under long days (16:8 h light/dark), whereas for all 13NH4+ influx experiments, northern analysis and competitive RT-PCR experiments, plants were grown vegetatively (8:16 h light/dark).

13N Influx Experiments

13NO3 was produced by proton bombardment of H2O by the cyclotron facility (TRIUMPH) of the University of British Columbia (Siddiqi et al., 1989). 13NO3 was reduced to 13NH4+ using Devarda's alloy, and 13NH3 was distilled into acidified nutrient solution (Wang et al., 1993). Plants used for influx experiments were grown in an open-top liquid culture system or in sterile media in 200-mL Magenta boxes (Magenta Corporation, Chicago) or 500-mL plastic growth containers (Sigma-Aldrich, St. Louis). Seeds were held at 4°C for 4 d in sterile dH2O and then seeded directly onto discs containing fine river sand (planting density, 1–3 seeds 1.5 cm−2) for the open system or onto a porous nylon mesh placed on floating discs (Sigma-Aldrich) for growth under sterile conditions. The discs were supported on polystyrene platforms, which floated in an 8-L basin. In the open system, plants were typically grown in dH2O for the first 10 d and then transferred to aerated complete nutrient solution (1 mm NH4NO3 except where indicated, 1 mm KH2PO4, 0.5 mm MgSO4, 0.25 mm CaSO4, 50 μm KCl, 25 μm H3BO3, 2.0 μm MnSO4·H2O, 2.0 μm ZnSO4·7H2O, 0.5 μm CuSO4·5H2O, 0.5 μm Na2MoO4, 20 μm Fe-EDTA, and 200 mg of CaCO3, pH 6.1) for 4 to 6 weeks. Nutrient solutions were replaced weekly (1 mm NH4NO3 plants) or daily (100 μm NH4NO3 plants). For plants grown under sterile conditions, the nutrient solution consisted of 1 mm KNO3 except where indicated, 29 mm Suc, 2 mm KH2PO4, 1.0 mm MgSO4, 1 mm CaCl2, 25 μm H3BO3, 2.0 μm MnSO4·H2O, 2.0 μm ZnSO4·7H2O, 0.5 μm CuSO4·5H2O, 0.5 μm Na2MoO4, 20 μm Fe-EDTA, and 200 mg L−1 CaCO3 at pH 6.1.

The influx experiments typically involved two pretreatments whereby 5-week-old plants were maintained for 4 more d on 1 mm NH4NO3, (N-sufficient plants) or on media without N (N-deprived plants). On the 4th d the plants were transferred to aerated uptake vessels for a 5-min pre-influx period in nonradioactive nutrient solution and then transferred to identical 13N-labeled NH4+ nutrient solution for 10 min followed by a 3-min desorption period in nonradioactive nutrient solution. Plant roots and/or shoots were subjected to a 15-s low speed centrifugation step and counted in a γ-counter (Minaxi, Auto-gamma 5000 series, Packard, Downer's Grove, IL). For determination of ammonium, nitrate, Gln pools, and %N and C, hydroponically grown plants were immersed in −N nutrient solution for 2 min, and then roots and shoots were frozen separately in liquid N2. Frozen tissues (17–20 plants per treatment) were ground in liquid N2 and lyophilized in a freeze drier for 4 d at −50°C. Three individual 20-mg fractions of dried tissue were extracted three times with 1 mL of 10 mm sodium acetate (pH 6.42), 4°C. Samples were centrifuged for 5 min at 14,000g (4°C) after each extraction. Pooled supernatant was subjected to three rounds of freeze (liquid N2) then thaw (iced water bath) periods and then centrifuged for 15 min at 14,000g (4°C). Samples were filtered (0.22 μm) and stored at −20°C. NO3 pools were estimated following the protocol of Cataldo et al. (1975). NH4+ and Gln pools were determined by derivatization of the extracts with AccQ (Waters, Milford, MA) followed by separation by HPLC (1090 LC, Hewlett-Packard, Palo Alto, CA) using a 4.6- × 25-mm C-18 column followed by fluorescent detection. Total N and C were measured from oven dried lyophilized tissues using an elemental analyzer (EA110 CHN-O, CE Instruments, Milan).

Isolation of amt1;1:T-DNA

The amt1;1:T-DNA line was identified from a pool of approximately 30,000 transfer DNA (T-DNA) lines composed of both the Feldmann CD5 (CD5 1–6; Forsthoefel et al., 1992) and “Jack” CD6 (CD6 1–6) series (Campisi et al., 1999). Extracted DNAs from subpools of these lines were obtained from the Arabidopsis Biological Resource Center (Ohio State University). The subpools of DNA from each set of lines were arranged in two multiplex arrays and screened using a PCR-based reverse genetic approach (Krysan et al., 1996) with forward (P1) and reverse (P2) oligonucleotides specific to AtAMT1;1. These oligonucleotides were used in conjunction with right (P3 and P4) and left (P5 and P6) border oligonucleotides of the T-DNA insertional elements of pD991 (Jack lines) and the 3850:1003 Ti plasmid (Feldman lines), respectively. Each 25-μL PCR contained 0.02 to 0.05 μg of pooled DNA, 25 pmol of each primer, 0.1 mm dNTPs, 2.5 μL of 10× PCR buffer, and 1 unit of Taq DNA polymerase (Invitrogen, Carlsbad, CA). PCR products were size-separated on 1% (w/v) agarose 1× Tris-acetate EDTA gels and blotted in 20× SSC onto nylon membrane (Hybond N+, Amersham Biosciences UK, Ltd., Little Chalfont, Buckinghamshire, UK; Sambrook et al., 1989). Blots were probed with a 1.7-kb random-primed [32P]dCTP labeled AtAMT1;1 cDNA followed by high-stringency washes (Rawat et al., 1999). PCR products that hybridized to the AtAMT1;1 cDNA were cloned into TOPO-TA (Invitrogen) and sequenced using Big Dye terminators (Applied Biosystems, Foster City, CA). A T-DNA insertion in AtAMT1;1 was identified in the Jack lines subpool CD6-4A. Twenty DNA subpools from CD6-4A (3,001–4,000) of 100 lines each (CD6-71 through CD6-90) were subjected to a PCR with primers P3 and P2 and Southern hybridization as described above. A positive signal in CD6-74 and CD6-85 was detected in the multiplex array of DNA pools. On the basis of this analysis, 3,350 was selected as the positive subpool. Approximately 300 seeds from subpool 3,350 were sown individually with a Pasteur pipette into separate pots with fine soil mix. Leaf tissue from 3-week-old seedlings was harvested for genomic DNA extraction according to McKinney et al. (1995). Using primers P2 and P3, a PCR was completed on each individual DNA sample, and amplified products were subjected to Southern hybridization with the AtAMT1;1 cDNA as described earlier. Three lines were identified yielding a positive signal for T-DNA insertion. The plants were grown to complete maturity, and the seeds were harvested. The Arabidopsis plants yielding a positive signal for a T-DNA insertion in AtAMT1;1 were backcrossed once to the Arabidopsis parental line (Col-3 gl1). Homozygous lines were then selected using the PCR with two sets of PCR primers. The first set (primers P3 and P7) were designed to identify the T-DNA insertion by amplifying between the right border of the T-DNA and the 3′ end of AtAMT1;1. The second set of primers (P8 and P7) were used to identify the non-tagged AtAMT1;1 allele by amplifying from −670 bp upstream of the first putative ATG through to the 3′ end of the of AtAMT1;1 gene. Putative homozygous lines were selected and analyzed further by Southern analysis and inverse PCR.

Southern Analysis and Amplification of Flanking T-DNA Genomic Sequences

Genomic DNA was isolated by cetyl-trimethyl-ammonium bromide extraction (Murray and Thomson, 1980) from both Col-3 gl1 and amt1;1:T-DNA leaves and digested with EcoRI and HindIII. Twenty micrograms of digested DNA was separated on a 0.8% (w/v) agarose 1× Tris-acetate EDTA gel and blotted onto nylon membranes in 20× SSC. The DNA was fixed to the membrane by baking at 120°C for 30 min. Blots were prehybridized in DIG-easy prehybridization mix (Roche Diagnostics, Indianapolis) at 42°C for 1 h and then incubated with PCR-amplified DIG-labeled uidA (GUS; accession no. A00196). The GUS cDNA was amplified from genomic DNA isolated from amt1;1:T-DNA using the PCR with primers P21 and P22. Amplified products were cloned into pGEMTeasy (Promega, Madison, WI) and sequenced. After hybridization, all membranes were washed twice for 15 min in 2× SSC, 1% (w/v) SDS at ambient temperature, twice at 68°C for 30 min in 0.1× SSC, 1% (w/v) SDS, and twice at ambient temperature in 0.1× SCC, 0.1% (w/v) SDS. Digoxygenin was detected using a commercial kit (Roche Diagnostics).

Inverse PCR was used to identify the nucleotide sequence adjacent to left border of the T-DNA insertion in amt1;1:T-DNA. Genomic DNA was isolated from amt1;1:T-DNA and digested with EcoRI. After digestion, the genomic DNA fragments were religated with T4 DNA ligase at 37°C for 2 h and then incubated at 16°C for a further 12 h. The ligated DNA was then subjected to a PCR using primers P23 and P24, which anneal in opposite directions to the left border of the T-DNA insert. Amplified products of the expected size were cloned into pGEMTeasy, and the cDNA insert was sequenced.

Northern Analysis

Total RNA was extracted from frozen plant tissues harvested from 4- to 6-week-old wild-type and amt1;1:T-DNA plants (roots and shoots) using the RNAeasy system (Qiagen USA, Valencia, CA). For northern analysis, 10 μg of total RNA per tissue was size separated on a 1× MOPS 1.2% (w/v) agarose gel containing formaldehyde (Sambrook et al., 1989) and blotted overnight onto Hybond N+ nylon membrane in 20× SSC. RNA was fixed to the membrane by baking at 120°C for 30 min. Blots were hybridized with full length DIG-labeled antisense AtAMT1;1 RNA produced using the SP6/T7 RNA DIG-labeling kit (Roche Diagnostics). Blots were hybridized overnight at 68°C in DIG-easy hybridization buffer (Roche Diagnostics). After hybridization, the blots were washed twice for 15 min in 2× SSC, 1% (w/v) SDS at ambient temperature, twice at 68°C for 30 min in 0.1× SSC, 1% (w/v) SDS, and twice for 15 min at ambient temperature in 0.1× SCC, 0.1% (w/v) SDS followed by detection of the digoxygenin label as previously described.

Competitive RT-PCR

Competitor RNA templates were prepared using the RT-PCR competitor construction kit (Ambion, Austin, TX). Each RNA competitor was transcribed from a modified deletion DNA template using T7 RNA polymerase. The AtAMT1;1 DNA template was constructed by cloning AtAMT1;1 (Rawat et al., 1999) into the NotI site of pSPORT2 (Invitrogen) followed by digestion with STYI, which removed 560 bp of the cDNA. The terminal STYI restriction sites of the gel extracted (QiaxII, Qiagen USA) pSPORT2/AtAMT1;1 deletion construct were ligated and the circularized plasmid amplified. The AtAMT2;1 and AtAMT1;3 deletion constructs were both prepared by first amplifying AtAMT2;1 and AtAMT1;3 cDNAs using RT-PCR on total RNA extracted from hydroponically grown Arabidopsis roots using PCR primers P9 and P10 for AtAMT2;1 and P11 and P12 for AtAMT1;3. Both PCR products were cloned into pGEMTeasy and sequenced. The 1.2-kb AtAMT2;1 fragment was inserted into the NOTI site of pSPORT2 and digested with STYI, which liberated a 658-bp fragment. The remaining pSPORT2/AtAMT2;1 construct was gel purified as above, and the terminal STYI sites were ligated. The pGEMTeasy/AtAMT1;3 construct was digested with AVAI, which liberated a 101-bp fragment. The remaining construct was gel extracted, and the terminal AVAI ends were ligated. For each of the above deletion constructs, a T7 RNA polymerase binding site was added to the 5′ end of the cDNA using the PCR and primer pairs (P13 and P14 for AtAMT1;1, P15 and P10 for AtAMT2;1, and P16 and P12 for AtAMT1;3). The AtAMT1;2 deletion template was synthesized using the PCR on AtAMT1;2 cDNA (Shelden et al., 2000) with primers P24 and P25, which deleted 435 bp from the 5′ end of AtAMT1;2 and added T7 RNA polymerase-binding site. PCR products were gel excised and purified (QiaxII, Qiagen USA). cRNA was generated using T7-RNA polymerase and RNase-resistant modified 2′-CTP (Ambion) in the presence of [32P]GTP. Products were size fractionated on a 4% (w/v) polyacrylamide 1× TBE gel containing 8 m urea. After autoradiography, the band of expected size was excised, and the RNase-resistant cRNA was eluted. The cRNA was quantified by measuring [32P]GTP incorporation using a scintillation counter (BeckmanCoulter, Fullerton, CA). Total RNA was extracted using the RNAeasy kit (Qiagen USA) from combined roots of six to eight individual plants per treatment. Total RNA was treated with DNase (DNase later, Ambion) and quantitated by UV spectroscopy. For the competitive RT-PCR experiments, one-step RT-PCR (Qiagen USA) was performed on total root RNA (25 ng) mixed with various concentrations of RNase-resistant competitor cRNA using primers P13 and P17 for AtAMT1;1, P26 and P18 for AtAMT1;2, P11 and P19 for AtAMT1;3, and P9 and P20 for AtAMT2;1. Amplified products were separated on agarose gels followed by staining with SYBR Gold (Molecular Dynamics, Sunnyvale, CA) and fluorescent signal immediately detected using a STORM imager (Amersham Biosciences).

Leaf Ultrastructure

Leaves of different ages were collected and sections sampled from the midway point between the tip of the leaf and the base of the petiole. Leaf samples were taken from plants grown either in the greenhouse in soil media with no supplementary lighting or in hydroponic tanks as described for the 13NH4+ influx experiments. Leaf samples that included the major vein were fixed overnight at 4°C in 2.5% (v/v) glutaraldehyde in 50 mm Na2HPO4, pH 7.0. Samples were washed and postfixed in 1% (v/v) OsO4 in 25 mm Na2HPO4 (pH 7.0) at room temperature for 1 h and then washed in 50 mm K2HPO4 (pH 7.0). Sections were dehydrated in successive ethanol washes (10%, 25%, 50%, 75%, 95%, and 100% [v/v]) and infiltrated and embedded in Spurr's resin at 60°C overnight. Sections (3 μm) were cut on a microtome (Leica Reichert Jung, Wetzlar, Germany) and stained with 0.5% (v/v) toluidine blue dissolved in 1% (w/v) sodium tetraborate (pH 9.5). Final sections were analyzed on a microscope (Zeiss, Welwyn Garden City, UK) and images captured using a digital camera (SPOT: Diagnostic Instruments, Sterling Heights, MI).

Statistical Analysis

Where reference to significant differences were made, analysis of variance of means was performed on the individual data points followed by two-tailed t tests (P < 0.05).

ACKNOWLEDGMENTS

We thank members of the A.D.M. Glass lab: Dr. Mamoru Okamoto, Anshuman Kumar, Dr. Manuela Simone, Dr. John J. Vidmar, and Aniko Varga for their participation in the 13NH4+ experiments. We also thank the particle acceleration facility “TRIUMPH” (Tri-university Meson Facility) at the University of British Columbia for the generation and supply of 13N. We also thank Joanna Maleszka (The Research School of Biological Sciences, The Australian National University) for the preparation of the leaf tissue sections and microscopy.

Footnotes

1This work was supported by the Natural Sciences and Engineering Research Council of Canada (grant to A.D.M.G. and postdoctoral fellowship to B.N.K.).

Article, publication date, and citation information can be found at www.plantphysiol.org/cgi/doi/10.1104/pp.102.010843.

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