Nitrate uptake

The first step in the assimilation of NO₃⁻ is usually uptake from the soil solution (reviewed by Glass and Siddiqi, 1995; Crawford and Glass, 1998; Forde and Clarkson, 1999; Galvan and Fernandez, 2001; Glass et al., 2001; Williams and Miller, 2001). Nitrate concentrations vary widely with typical levels in agricultural soils ranging from 1–10 mM. Uptake occurs in the outer cell layers of the root by active transport processes mediated by H+ cotransport. In other words, the energy stored in the plasma membrane H+ gradient drives NO₃⁻ transport into epidermal and cortical cells. Such activity can be followed by measuring the depletion of NO₃⁻ from the media surrounding roots or by monitoring the changes in electrical potential across the plasma membrane with microelectrodes as NO₃⁻ uptake coincides with a depolarization of the plasma membrane, i.e. becomes more positive inside (Miller et al., 2001). An excellent and extensive electrophysiological study using Arabidopsis root hairs has been reported (Meharg and Blatt, 1995). Such experiments uncovered two general systems that account for NO₃⁻ uptake: a high affinity system, whose K_m for NO₃⁻ ranges from 10−100 µM, and a low affinity system, which typically shows linear or nonsaturable kinetics and is most evident above 500 µM NO₃⁻. Both systems show depolarization in response to NO₃⁻ indicating that both are active and involve cotransport. The high and low affinity systems correspond to the two classical mechanisms, Mechanism I and Mechanism II, respectively, described by Epstein, who first showed ion uptake in plant roots obeyed Michaelis-Menten kinetics (Epstein, 1953; Epstein, 1972).

An important aspect of NO₃⁻ uptake is that it is regulated (reviewed by Crawford and Glass, 1998; Daniel-Vedele et al., 1998; Forde and Clarkson, 1999; Forde, 2000; Galvan and Fernandez, 2001; Glass et al., 2001; Williams and Miller, 2001). Various environmental and internal signals influence the rate of uptake into roots. Nitrate itself acts as an inducer. When plants are deprived of NO₃⁻ (either by starving them of nitrogen or by providing them with another form of nitrogen such as NH₄⁺ or an amino acid), they retain a basal or constitutive level of NO₃⁻ uptake that has both high and low affinity components (abbreviated as cHATS and cLATS, respectively). Upon exposure to NO₃⁻, uptake rates increase. The characteristics of such induced activity (abbreviated iHATS) indicate that it is a high affinity system distinct from cHATS. In addition to NO₃⁻ induction, uptake shows feedback inhibition and responds to signals from the shoot. Nitrate uptake decreases when plants are fed NH₄⁺ or high levels of NO₃⁻. Uptake also responds to circadian rhythms, reduced carbon, and shoot nitrogen demand. A classic example of shoot demand comes from split root experiments where one half of the roots from a single plant are placed in one chamber and the other half are placed in a second chamber (Figure 2). The first half is monitored for NO₃⁻ uptake in a solution with ample NO₃⁻. If the remaining half of the roots is placed in a solution with low NO₃⁻, uptake rates increase in the first half. It is thought that the shoots sense a reduction in overall nitrogen availability when the second group of roots is placed in low NO₃⁻ solutions. The shoots then respond by sending a signal to the roots to increase NO₃⁻ uptake.

Nitrate reduction

Once NO₃⁻ has been taken up into the cytosol, it can be (1) mobilized into the xylem for transport to the shoot, (2) transported into the vacuole for storage or (3) reduced to nitrite, a reaction that is catalyzed by NO₃⁻ reductase (NR), a cytosolic enzyme that uses NAD(P)H for reductant (Campbell, 1999; Campbell, 2001). Nitrate reduction is the first committed step in the NO₃⁻ assimilation pathway and is regulated at both transcriptional and post-translational levels. Both the NO₃⁻ reductase genes (NIA) and the NR protein respond to many signals (Crawford, 1995; Kaiser et al., 1999; Stitt, 1999; Tischner, 2000; MacKintosh and Meek, 2001). As is the case for NO₃⁻ uptake, NO₃⁻ serves a key inducer that rapidly activates transcription of the NIA genes. Sucrose, light and cytokinin enhance NIA induction while NH₄⁺ and amino acids repress expression. NIA genes also respond to circadian rhythms.

Another well understood mechanism of NR control occurs at the protein level (Kaiser et al., 1999; MacKintosh and Meek, 2001). NR produces nitrite, which is toxic to plants if it accumulates. Nitrite is transported into the chloroplasts in green tissues (plastids in non-green tissues), where it is reduced to NH₄⁺ by nitrite reductase (NiR) using reduced ferredoxin. If nitrite reduction is slowed (e.g. caused by a reduction in light, reduced carbon or oxygen), nitrite levels could increase to toxic levels if NR activity is not inhibited. Plants have a rapid and reversible mechanism for doing just this. NR is phosphorylated by one of several kinases on a key serine in the first hinge region, which separates the Mo-cofactor and heme-binding regions of NR. NR has three redox centers (FAD, heme and the Mo-cofactor) that are bound by three distinct regions of the protein (Campbell, 1999; Campbell, 2001). The Mo-cofactor transfers electrons to NO₃⁻ while the heme shuttles electrons from a FAD prosthetic group to the Mo-factor. If NR is phosphorylated in the hinge 1 region, it can bind a 14-3-3 dimer, which in turn inactivates the enzyme. Removal of the 14-3-3 dimer or dephosphorylation of NR by a phosphatase results in reactivation of the enzyme. Exposure of plant to light, high CO₂ and oxygen levels leads to most of the NR being active while darkness, low CO₂ levels and hypoxia result in most of the NR being inactive.

Nitrate assimilatory mutants of Arabidopsis

In the NO₃⁻ assimilation field, Arabidopsis was the first plant used for mutant isolation. In the early seventies, a graduate student Fietje Braaksma isolated chlorate resistant mutants (chl) of Arabidopsis in the lab of W.J. Feenstra (Braaksma and Feenstra, 1973). Chlorate is the chlorine analog of NO₃⁻, which, when taken up and reduced to chlorite, is toxic to plants. Chlorate resistant mutants had been identified in fungi and were shown to be defective in NO₃⁻ uptake, reduction and regulation (Cove, 1979; Marzluf, 1981; Crawford and Arst, 1993), but no such mutants had been described in plants. In her initial report, Braaksma described three mutants, two of which were allelic. B1 and B3 (now called chl1 or nrt1.1) had normal levels of NR activity but reduced levels of chlorine accumulation (a measure of ClO₃⁻ uptake). B2-1 (now called chl2) had much reduced NR activity. These data uncovered two genes, one involved in NO₃⁻ reduction and another in chlorate uptake. After Braaksma and Feenstra's initial publication, additional chl mutants were described in Arabidopsis (Braaksma and Feenstra, 1982; Braaksma and Feenstra, 1982), in cells lines of Nicotiana (Muller and Grafe, 1978) and in barley plants (Kleinhofs et al., 1978). As these and other mutants were characterised in angiosperms, they were all found to fall into two classes: those having mutations in NIA (R) genes and those having mutations in CNX genes needed for the synthesis of the Mo-cofactor (Crawford, 1992; Warner and Kleinhofs, 1992; Crawford and Arst, 1993; Hoff et al., 1994; Mendel, 1997). Both classes have reduced NR activity. No mutations were found in NO₃⁻ uptake or regulatory genes except for chl1.

Further work on the molecular genetics of NO₃⁻ reduction in Arabidopsis revealed that it had two NIA genes and several CNX genes (Crawford, 1992). The NIA2 gene, which encodes for most of the NR activity in the plant (85–90%), corresponds to the chl3 locus at position 56 in the middle of chromosome 1 (Crawford et al., 1988; Wilkinson and Crawford, 1991). The NIA1 gene encodes for 10–15% of the NR activity and is located near the end of chromosome 1 at position 116 near the GL2 gene (Cheng et al., 1988; Wilkinson and Crawford, 1993). Both NIA genes are regulated by NO₃⁻ and circadian rhythms and are expressed in leaves and roots (Cheng et al., 1988; Crawford et al., 1988; Cheng et al., 1991; Pilgrim et al., 1993). Such duplication of genes is found over much of the Arabidopsis genome (Blanc et al., 2000), and the NIA genes provided an early example of this. It is interesting that even though there is genetic redundancy, one can still isolate chlorate resistant mutants in which only the NIA2 gene is inactivated. This is because the two genes have diverged enough so that NIA2 accounts for most of the NR activity. NIA2 (chl3) mutants still show some chlorate sensitivity due to NIA1, but they are much less sensitive than wildtype plants. The residual NR activity (10–15% wildtype) from the NIA1 gene is sufficient for normal growth on NO₃⁻ as the sole nitrogen source (Crawford, 1995). For such a key enzyme, this result was surprising but was found to be true in other plants as well (reviewed in (Crawford, 1995)). When it was discovered that NR is post-translationally regulated, an explanation for this puzzling result arose. In wildtype tobacco plants, the NR is not 100% active when measured in the presence of magnesium but oscillates between 20–90% of full potential activity, which is measured in the presence of EDTA (Scheible et al., 1997a). In NR mutants, the extent of inactivation is reduced so that the reduced NR levels in the mutant are compensated by less inactivation of NR (Scheible et al., 1997a). It is not until NR activity levels drop below 10% of wildtype levels that one observes nitrogen deficiencies in the mutants. In Arabidopsis, a nia1nia2 double mutant has been described with a deletion of the NIA2 gene and a point mutation in the NIA1 gene that has only 1% of wildtype level of activity (Wilkinson and Crawford, 1993). This mutant has grows poorly on NO₃⁻ alone but does show some growth.

The other class of NR-deficient mutants is cnx. Four cnx mutants have been identified in Arabidopsis: chl2, chl4 (B25), chl6 (B73) and chl7 (Crawford, 1992). An interesting feature of these mutants is that three of them (chl2, chl6 and chl7) are unable to grow on 0.1 mM tungstate, a useful tool for testing the loss of Mo-cofactor (LaBrie et al., 1992). Further work has lead to the identification of six genes required for synthesis of the Mo-cofactor in Arabidopsis (Mendel, 1997) (Figure 3). The early steps convert a guanosine nucleotide into an intermediate called Precursor Z. These steps are mediated by the products of the CNX2 and CNX3 genes (Hoff et al., 1995). Precursor Z is converted into molybdopterin by the CNX5, CNX6 and CNX7 products (Mendel, 1997). Molybdate is then added to molybdopterin by the two domain protein CNX1, which is homologous to the mammalian neuroprotein Gephyrin and binds the cell's cytoskeleton as does Gephyrin (Stallmeyer et al., 1995; Kuper et al., 2000; Schwarz et al., 2000). The CNX1 gene is the same as CHL6 (originally named B73) (Schwarz et al., 2000).

The NRT1 family of nitrate transporters

Unlike NO₃⁻ reduction, the molecular genetics of NO₃⁻ uptake had been hampered by the fact that no uptake mutants were known in plants for sometime except for chl1 in Arabidopsis (in the new nomenclature the gene is called AtNRT1.1). Ironically, this was the first chlorate-resistant mutant ever identified in plants, and mutations at this locus are found more frequently than in any other gene in chlorate selections. The original work on chl1 indicated that it was strongly inhibited in chlorate uptake and partially defective in NO₃⁻ uptake (Doddema et al., 1978). The latter defect appeared to be a reduction in low affinity but not high affinity NO₃⁻ uptake activity (Doddema and Telkamp, 1979). Subsequent cloning of the gene from a T-DNA-tagged line revealed that it encoded a hydrophobic protein with twelve membrane spanning regions indicative of a cotransporter (Tsay et al., 1993a). Expression of the CHL1 protein in Xenopus oocytes showed it was a NO₃⁻ transporter (Tsay et al., 1993a) with, surprisingly, biphasic kinetics (Huang et al., 1996; Liu et al., 1999). These experiments along with studies of NO₃⁻ uptake in whole plants demonstrated that CHL1 is a dual affinity transporter contributing to both HATS and LATS (Huang et al., 1996; Touraine and Glass, 1997; Wang et al., 1998; Liu et al., 1999). CHL1's contribution, however, depends on the environmental conditions used for plant growth. One key effector is NH₄⁺. Overall NO₃⁻ uptake is reduced when plants are grown with NH₄NO₃ as compared to KNO3; however, CHL1's contribution is the greatest when plants are grown with NH₄⁺; that is, chl1 mutants show the largest reduction in uptake when grown with NH₄⁺ than without (Huang et al., 1996; Touraine and Glass, 1997). Another critical parameter is pH. CHL1 is more highly expressed at acidic pHs (Tsay et al., 1993a) and makes a greater contribution to cHATS at pH 5.5 than pH 7.0 (Wang et al., 1998). CHL1 expression also shows diurnal variation, and the reduction in the dark can be reversed by supplying sucrose indicating that carbon signals affect CHL1 (Lejay et al., 1999). Nitrogen metabolites, however, do not appear to repress CHL1 as depriving Arabidopsis of nitrogen does not increase CHL1 expression (Lejay et al., 1999).

Recent experiments have shown that CHL1 expression is targeted to actively growing regions of roots and shoots (Guo et al., 2001). High levels of expression are found in root tips, lateral root primordia, young leaves and flower buds. Little expression over background is found in mature leaves, flowers and the apical meristem. In the mature parts of the root, intermediate expression is found in the stele and little in the outer layers of the root. chl1 mutants also are defective in growth of nascent organs. Thus, the function of CHL1 is not simply to take up nitrate from the soil by the main body of the root but also to support the growth of nascent and actively growing organs in both roots and shoots.

After cloning CHL1, four additional genes with similar sequences were found in Arabidopsis (Table 1). AtNRT1.2 (originally named NTL1) was cloned by homology to CHL1 and found to be 36% identical at the amino acid level (Huang et al., 1999). Further work showed that it is a low affinity NO₃⁻ transporter (K_m of 6 mM in oocytes) that is constitutively expressed in root epidermal and root hair cells. This finding indicates that AtNRT1.2 is a component of cLATS. A second gene AtPTR2B encodes a peptide transporter (Rentsch et al., 1995; Song et al., 1996). In fact, almost all the CHL1-related genes which have been described from bacteria, yeast and mammals have been identified as transporters of small oligopeptides including di- and tri-peptides (Steiner et al., 1995; Graul and Sadee, 1997; Crawford and Glass, 1998). Why transporters with specificity to NO₃⁻ should be related to peptide transporters is not clear. A most interesting protein, NRT1.2 from Brassica napus, transports histidine and NO₃⁻ (Zhou et al., 1998), indicating that this protein retains substrate specificities found in all members of the family. This protein is also the most similar to CHL1 being 91% identical. The entire gene family has been named PRT for peptide transporter and POT for H+ -dependent oligopeptide transporter. This family was tentatively placed in the Major Facilitator Superfamily (MFS) of transporters (Pao et al., 1998) but now is its own family (Saier, 2000). The PTR and POT names are misleading as they do not include the NO₃⁻ transporters in the family. Two other genes in this family have been cloned from Arabidopsis and named NTP2 (now called AtNRT1.4) and NTP3 (now called AtNRT1.3) (Hatzfeld and Saito, 1999), but their functions are not known, and they do not appear to be nitrate-regulated (Glass et al., 2001).

The NRT2 family of transporters

The second family of NO₃⁻ transporter genes (NRT2) was first discovered in Aspergillus. A chlorate-resistant mutant called crnA was identified with a defect in NO₃⁻ uptake (Brownlee and Arst, 1983). It was subsequently cloned and shown to be NO₃⁻—regulated and NH₄⁺-repressed and to encode a transport protein (Unkles et al., 1991). Subsequently, two related genes (NAR-3 and NAR-4) were isolated from Chlamydomonas (Quesada et al., 1994). NAR-3 (CrNrt2.1) was shown to encode a high affinity, bi-specific NO₃⁻/NO₂⁻ transporter while NAR-4 (CrNrt2.2) encodes a high affinity NO₃⁻—specific transporter (Galvan and Fernandez, 2001). The NAR-3 protein is about 30% identical to CRNA. Independently, a barley cDNA clone (BCH1) was identified by PCR amplification using degenerate primers corresponding to two conserved motifs found in a small group of MFS transporters that included CRNA (Trueman et al., 1996; Trueman et al., 1996). BCH1 (now called HvNRT2.1) is 32% identical to CRNA and 50% identical to CrNRT2.1. Subsequently, NRT2 genes have been identified in six plant species as well as Chlamydomonas, Aspergillus, Hansenula and E. coli (reviewed by Crawford and Glass, 1998; Daniel-Vedele et al., 1998; Forde, 2000; Galvan and Fernandez, 2001; Glass et al., 2001; Williams and Miller, 2001). These genes are found in the NO₃⁻/NO₂⁻ porter (NNP) family within the major facilitator superfamily (Saier, 2000).

In Arabidopsis a total of 7 NRT2 genes have been found (Table 2) (Forde, 2000; Glass et al., 2001; Okamoto et al., 2001). The first two identified were AtNRT2.1 and AtNRT2.2 ((Filleur and Daniel-Vedele, 1999; Zhuo et al., 1999) and Genbank Accession # AF019748 and Z97058). These genes are about 1 kb apart in a tail to tail orientation on chromosome 1 at position 10–12 cM, not far from CHL1. NRT2.1 shows strong regulated expression in roots (Filleur and Daniel-Vedele, 1999; Lejay et al., 1999; Zhuo et al., 1999; Cerezo et al., 2001) while little mRNA is detected for NRT2.2 (Zhuo et al., 1999). NRT2.1 is induced by NO₃⁻ or by nitrogen starvation and is repressed by NH₄⁺, amino acids (especially arg) and by growth on high levels of NO₃⁻. These properties are all characteristic of the inducible HATS system, which is NO₃⁻—inducible and feedback repressed by nitrogen metabolites or high NO₃⁻ levels. NRT2 has been proposed to be a component of the iHATS in plants (reviewed by (Forde, 2000)).

Direct demonstration that NRT2.1 is component of iHATS has come from mutant and heterologous expression studies. Arabidopsis transgenic lines containing antisense constructs of AtNRT2.1 (Forde, 2000) and an Arabidopsis deletion mutant removing all of NRT2.1 and part of NRT2.2 (Filleur et al., 2001) have been constructed and shown to be defective in HATS but not LATS. Furthermore, the NRT2.1/2.2 mutant is defective in iHATS activities (i.e. in NO₃⁻—inducible, starvation-derepressible and NH₄⁺-repressible high affinity uptake) (Cerezo et al., 2001). The CRNA protein has been functionally expressed in Xenopus oocytes and found to have two transport mechanisms: a H+ -coupled mechanism with a K_m for NO₃⁻ of 24 µM and a passive mechanism with a K_m for NO₃⁻ of 126 µM (Zhou et al., 2000b). It was suggested that under conditions of high external NO₃⁻ concentrations the passive mechanism would provide an energetic advantage to the fungus by enabling it to absorb NO₃⁻ without the energetically expensive coupling to H+ fluxes. Whether plant NRT2 transporters function in similar ways is still unknown. The Chlamydomonas NRT2.1 protein expresses high affinity NO₃⁻ transport activity in Xenopus oocytes but only if co-expressed the a second protein (NAR2), indicating that this system requires two components (Zhou et al., 2000a).

Overview of nitrate transporter regulation and function

Additional aspects of overall transporter regulation have emerged from Arabidopsis. Both CHL1 and NRT2.1 display diurnal rhythms having low levels of mRNA at night (Lejay et al., 1999). The low levels at night can be enhanced by the addition of sucrose. These two genes are also NO₃⁻—responsive but CHL1 does not display the feedback regulation that NRT2.1 does as it is not repressed by NH₄⁺ treatment nor induced by nitrogen starvation (Filleur and Daniel-Vedele, 1999; Lejay et al., 1999). CHL1 does respond to pH as its mRNA levels are enhanced at more acidic pH (Tsay et al., 1993a).

With all this information, it has become possible to associate a given gene with a NO₃⁻ uptake system. The two genes for which we have the most physiological data are CHL1 and NRT2.1. Present data indicate that NRT2.1 is a major component of iHATS that would be especially significant when plants are provided low levels or are deprived of nitrogen. The role of CHL1 is more complicated. The expression patterns of this gene and the properties of the CHL1 protein suggest that it could contribute to all three uptake systems in that it is expressed in the absence of NO₃⁻, its expression is enhanced by NO₃⁻ and it has dual affinity uptake kinetics. Measurements of uptake in whole plants have indicated that CHL1 does contribute to cHATS and LATS, but its role in iHATS is controversial (Wang et al., 1998; Liu et al., 1999; Cerezo et al., 2001; Glass et al., 2001). These experiments have also indicated that CHL1's contribution depends heavily on the presence of NH₄⁺, which would also repress NRT2.1 expression. In the presence of NH₄⁺, chl1 mutants have significantly reduced NO₃⁻ uptake activity compared with wildtype plants for both cHATS and LATS. This is true for plants grown with or without NO₃⁻. Thus, unlike NRT2.1, it is difficult to assign CHL1 to one particular system. It is also appears that cHATS, iHATS and LATS are not static but dynamic systems whose composition changes depending on the environmental conditions in which it is grown and on the developmental stage of the plant. Other Arabidopsis transporters that are known to contribute to the NO₃⁻ uptake systems are NRT2.1, a component of iHATS (Filleur and Daniel-Vedele, 1999; Cerezo et al., 2001; Filleur et al., 2001), NRT1.2, whose properties indicate it is a constitutive component of LATS (Huang et al., 1999), and CHL8, whose gene has not been cloned but which contributes to cHATS (Wang and Crawford, 1996).

Nitrate regulation of gene expression

As can be seen from the above discussion, NO₃⁻ is not only a substrate for the NO₃⁻ assimilatory pathway, but it is also a key signal. One of the most studied aspects of NO₃⁻ signalling is induction of gene expression (reviewed by (Redinbaugh and Campbell, 1991; Crawford, 1995; Koch, 1997; Stitt, 1999; Tischner, 2000; Wang et al., 2000; Coruzzi and Bush, 2001). Many genes involved in different aspects of nitrogen and carbon metabolism are induced by NO₃⁻. This effect can be observed by treating plants that are either starved for nitrogen or grown with a reduced source of nitrogen such as NH₄⁺ or amino acids. Genes that respond to NO₃⁻ have been grouped based on their metabolic niche as follows: NO₃⁻ assimilation, NH₄⁺ assimilation, ferredoxin reduction, oxidative pentose phosphate pathway and organic acid synthesis. Nitrate also mobilises carbon into organic acids by suppressing starch synthesis.

The primary NO₃⁻ assimilatory genes (NIA, NII and NRT) all respond to NO₃⁻ (Tang and Wu, 1957; Cheng et al., 1986; Crawford et al., 1986; Melzer et al., 1989; Cheng et al., 1991; Gowri et al., 1992; Aslam et al., 1993; Tischner et al., 1993; Koch, 1997; Stitt, 1999; Tischner, 2000). The NIA genes have been the most studied and respond within minutes to NO₃⁻ at concentrations as low as 10 µM. In addition to NII, there are a group of responsive genes that contribute to nitrite reduction by providing reductant (encoding ferredoxin (Fd), ferredoxin NADP+ oxidoreductase (FNR), 6-phosphogluconate dehydrogenase (6PGDH) and 6-phosphogluconate dehydrogenase (6PGDH) and S-adenosyl-L-methionine-dependent uroporphyrinogen III methyltransferase (UPM1), which makes a prosthetic group for the NiR enzyme) (reviewed by (Stitt, 1999; Wang et al., 2000)). Genes involved in NH₄⁺ assimilation, encoding specific isoforms of glutamine synthetase and glutamate synthase, are also induced (reviewed by (Lam et al., 1996; Koch, 1997; Stitt, 1999)). Responsive genes involved in starch and organic acid metabolism encode phosphoenolpyruvate carboxylase (PEPC, involved in organic acid metabolism), which is induced, and ADP-glucose pyrophosphorylase (agpS2, involved in starch synthesis), which is repressed by NO₃⁻ (Scheible et al., 1997b). Other genes that encode organic acid metabolic enzymes—cytosolic pyruvate kinase (PKc), citrate synthase (CS), and NADP-isocitrate dehydrogenase (ICDH1)—respond to NO₃⁻ (Scheible et al., 1997bb).

A microarray analysis of NO₃⁻—responsive genes has been reported for Arabidopsis (Wang et al., 2000). Using glass slides with approximately 5500 Arabidopsis genes/clones, mRNA from plants treated with low NO₃⁻ (250 µM) for short times (20 min) and with high NO₃⁻ (5–10 mM) for longer times (2 hrs) was analysed to detect both rapid response genes and responsive genes in general. Besides detecting many of the genes described above, new responsive genes were identified. Two genes of the nonoxidative branch of the pentose phosphate pathway—transketolase and transaldolase—were induced. Other metabolic genes that were induced included malate dehydrogenase, asparagine synthetase, histidine decarboxylase and two methyltransferases. Several other very interesting genes were induced including a myb transcription factor gene, a tonoplast high capacity calcium antiporter gene CAX1, the senescence-associated gene SAG21 and the nonsymbiotic hemoglobin gene AHB1. Several genes were found to be repressed by NO₃⁻ including the NH₄⁺ transporter gene AMT1;1. The general conclusions that came from this and other studies are the following. First, the most responsive genes are those that are involved directly or indirectly with nitrite reduction (encoding NiR, Fd, FNR, UMP1, G6PDH, 6PGDH, transketolase and transaldolase). Nitrite is toxic to plants so that it makes sense that the genes critical for nitrite reduction would be rapidly induced when NO₃⁻ is present. Second, different genes show different response patterns indicating that multiple signalling and regulatory events are occurring as plants take up and assimilate NO₃⁻. For example, some genes show reduced expression after 2 hrs in low NO₃⁻ while others do not. Key signals that may be dictating expression levels are NO₃⁻, reduced nitrogen and carbon metabolites. As NO₃⁻ gets depleted by assimilation or is accumulated in the vacuole, some genes will sense the drop in NO₃⁻ and begin to diminish their expression. In addition, the products of NO₃⁻ assimilation including NH₄⁺ and amino acids, especially glutamine and glutamate, are also thought to feedback repress genes in the pathway (Hoff et al., 1994; Dzuibany et al., 1998; Tischner, 2000). Carbon metabolites sucrose (Cheng et al., 1992) and malate (Muller et al., 2001) also have been shown to serve as signals.

With such elaborate signalling, it is clear that regulatory networks exist to control NO₃⁻ assimilation and integrate it with nitrogen and carbon metabolism. Our understanding of these networks is in its infancy. The discussion below is restricted to NO₃⁻ signalling. Several promoter elements have been identified that are needed for NO₃⁻ induction of NiR (Rastogi et al., 1993; Rastogi et al., 1997) and NR (Lin et al., 1994; Hwang et al., 1997) genes in tobacco (reviewed by Rothstein and Sivasankar, 1999). With the use of inhibitors, it has been shown that NO₃⁻ induction is a primary response (i.e. not blocked by protein synthesis inhibitors (Gowri et al., 1992; Redinbaugh and Campbell, 1993; Ritchie et al., 1994; Redinbaugh and Campbell, 1998)) and involves calcium (Sakakibara et al., 1996; Sakakibara et al., 1997; Sueyoshi et al., 1999) as well as kinases and phosphatases (Champigny and Foyer, 1992; Sakakibara et al., 1997; Sueyoshi et al., 1999). There have been two reported attempts to find NO₃⁻ regulatory mutants in Arabidopsis. The first selected for mutants that were resistant to chlorate after a transient treatment with low NO₃⁻ (Lin and Cheng, 1997). A mutant (cr88) was found that was defective in light reception and was designated as a hy mutant (Lin and Cheng, 1997; Cao et al., 2000). The other was a screen using transgenic plants containing the tobacco NiR promoter fused to GUS (Leydecker et al., 2000). This screen produced cnx mutants that overproduced NiR mRNA in the absence of NO₃⁻. Thus, no regulatory mutants defective in NO₃⁻ induction have been reported. However, a NO₃⁻—regulated gene (ANR1) has been identified which encodes a MADS box transcription factor that is involved in nitrate regulation of root development. This gene is discussed in more detail below (Zhang and Forde, 1998).

Kinetics and Energetics of NH₄⁺ Uptake

In common with other plants, the kinetics of NH₄⁺ uptake in Arabidopsis are biphasic, being separable into a high-affinity system (HATS) and a low-affinity system (LATS) (Rawat et al., 1999). The LATS only becomes evident at external NH₄⁺ concentrations above 1 mM (Rawat et al., 1999). The high-affinity system displays typical Michaelis-Menten kinetics (Rawat et al., 1999; Shelden et al., 2001), while the LATS has linear (non-saturable) kinetics (Rawat et al., 1999), consistent with it being mediated by a passive transport system such as a channel (see below). The affinity of the HATS for its substrate, as estimated from ¹³NH₄⁺ labelling studies, falls within the range reported in other plants (Rawat et al., 1999). For seedlings maintained on NH4NO3, the estimated K_m values for NH₄⁺ were from 85 to 168 µM, compared with values in the range 20–360 µM for other plant species (von Wiren et al., 2001). Using an electrophysiological method, a similar K_mA0value of 46 ± 5 µM was obtained for NH4NO₃⁻grown Arabidopsis seedlings, but seedlings that had been N-starved or grown on NO₃⁻ alone for 3 d gave K_m values of 7–8 µM (Shelden et al., 2001).

It is now generally accepted that NH₄⁺, rather than the unprotonated species NH₃, is the transported form (Forde and Clarkson, 1999; von Wiren et al., 2001). However the precise mechanism of NH₄⁺ influx across the plasma membrane is still not fully resolved. Based on typical estimates for cytosolic NH₄⁺ concentrations and the electrical potential across the plasma membrane, thermodynamic considerations imply that in the range of external NH₄⁺ concentrations that roots are exposed to in well-aerated soils (i.e. <50 µM) the uptake of NH₄⁺ will require the input of energy (Forde and Clarkson, 1999). This could be achieved, for example, by a high-affinity H+/NH₄⁺ co-transporter that coupled NH₄⁺ influx to the H+ gradient generated by the plasma membrane H+ pump. The importance of the H+ gradient in driving NH₄⁺ influx by the HATS in rice roots has been demonstrated using protonophores and ATPase inhibitors (Wang et al., 1993), but direct evidence for H+/NH₄⁺ symport in plant roots is still lacking. In the bacterium Corynebacterium glutamicum, the amt NH₄⁺ transporter (which is homologous to higher plant NH₄⁺ transporters) has been proposed to operate via an NH₄⁺ uniport mechanism that is driven by the membrane potential (Siewe et al., 1996).

The net uptake of any ion is the result of the opposing processes of influx and efflux, and under some conditions it is known that NH₄⁺ efflux can have a significant influence on net NH₄⁺ uptake (Forde and Clarkson, 1999). Rates of NH₄⁺ efflux in Arabidopsis (Rawat et al., 1999) and in other plants (Wang et al., 1993; Kronzucker et al., 1995) are positively correlated with the cytosolic NH₄⁺ concentration. When Arabidopsis seedlings were grown on NH₄⁺-rich medium (1 mM NH₄⁺) the rate of NH₄⁺ efflux was estimated to be as much as 35% of the influx rate (Rawat et al., 1999). As discussed elsewhere (Forde and Clarkson, 1999), because the efflux of the protonated form of NH₄⁺ is likely to be thermodynamically uphill, it is thought that it is actually NH₃ that diffuses passively from the cells and that this is then protonated in the external solution giving the false impression that the ionic form had crossed the plasma membrane (Figure 4).

Regulation of NH4+ Uptake: the Physiological Evidence

It has been established for a wide range of plant species that the HATS for NH₄⁺ (but not the LATS) is highly regulated according to the N nutrition of the plant (Forde and Clarkson, 1999; von Wiren et al., 2001). A number of studies have now confirmed that the same is true in Arabidopsis (Gazzarrini et al., 1999; Rawat et al., 1999; Shelden et al., 2001). For example, transferring Arabidopsis plants from high NH4NO3 concentrations to low led to a 12-fold increase in HATS activity (Rawat et al., 1999), and when N-depleted plants were resupplied with NH4NO3, the NH₄⁺ influx rate declined rapidly (Rawat et al., 1999). In the same experiments, the LATS activity was independent of the external NH₄⁺ supply and the plant's N status.

As already noted above, the kinetic properties of the HATS can also be affected by changes in N nutrition. In rice, a positive correlation was found between the external NH₄⁺ supply and the affinity of the HATS for NH₄⁺ (Wang et al., 1993). While Rawat et al., working with Arabidopsis, found no simple relationship between the ambient N supply and the affinity of the HATS for NH₄⁺ (Rawat et al., 1999), a more recent study reported that when Arabidopsis seedlings were grown for 3 d in the absence of NH₄⁺ (whether N-deprived or with NO₃⁻ as sole N source) the K_m for NH₄⁺ declined from 46 µM to <10 µM (Shelden et al., 2001). A possibly critical difference between the two studies is that in the former case the ‘low N’ plants were grown on 0.1 mM NH4NO3 and so were not fully NH₄⁺-deprived. Perhaps the very high-affinity HATS is synthesised or activated only when the external NH₄⁺ concentration is very low or zero. NH₄⁺ influx in Arabidopsis roots is diurnally regulated, reaching a maximum at the end of the light period and declining markedly with the onset of darkness (Gazzarrini et al., 1999). This suggests that the C status of the plant may have a regulatory effect on HATS activity that is somehow transmitted from shoot to root.

Structure and Biochemical Properties of the AMT Family of NH₄⁺ Transporters

The Arabidopsis AtAMT1.1 gene was the first NH₄⁺ transporter gene to be cloned from higher plants (Ninnemann et al., 1994). Subsequently, two closely related cDNAs (AtAMT1.2 and AtAMT1.3) were isolated by homology and shown to be able, like AtAMT1.1, to complement an NH₄⁺ transport mutant of yeast (Gazzarrini et al., 1999). The completed Arabidopsis genome sequence reveals that there are two further members of the AMT1 family (AtAMT1.4 and AtAMT1.5) (Table 3), but no further data are available on these. A sixth gene (AtAMT2), which is more distantly related to the other AMT genes, has been identified and partially characterised (Sohlenkamp et al., 2000).

The AMT genes encode hydrophobic proteins of 475–514 amino acids and belong to a probably ubiquitous gene family (the AMT/MEP family), which has known members in bacteria, archaea, fungi, plants and animals (Saier et al., 1999). All functionally characterised members of the family catalyse NH₄⁺ uptake and most also transport the NH₄⁺ analogue, methylammonium. Figure 5 illustrates the phylogenetic relationships between the Arabidopsis AMT transporters and selected members of the wider AMT/MEP family. The AtAMT1 genes seem to fall into three subfamilies, one that includes AtAMT1.1, AtAMT1.3 and AtAMT1.5 (the AtAMT1.1 subfamily), and two others that have single members, AtAMT1.2 and AtAMT1.4. The tomato LeAMT1.1 and the Lotus japonicus LjAMT1.1 genes belong to the AtAMT1.1 subfamily, while the tomato LeAMT1.2 gene is most closely related to AtAMT1.2. The close relationship between AtAMT1.3 and AtAMT1.5, together with their occurrence adjacent to each other within a 6 kb region of chromosome 3, suggests that they represent a relatively recent gene duplication. The AtAMT2 gene is the most divergent of the Arabidopsis AMT sequences, being more closely related to the yeast and bacterial and yeast AMT/MEP genes than to the plant ones (Figure 5).

The tomato LeAMT1.3 transporter has an N-terminus that is 20 amino acids shorter than the other AMT1 polypeptides and two untranslated open reading frames are found in its 5′-untranslated region, something often associated with regulatory genes (von Wiren et al., 2000b). Surprisingly, there appears to be no direct ortholog of LeAMT1.3 in the Arabidopsis genome.

Computer-based predictions that the E. coli AMTB protein has 12 transmembrane domains, with the N- and C-termini located on the interior face of the membrane, were recently confirmed using fusions with marker proteins (Thomas et al., 2000). However, sequence alignments indicate that the most N-terminal of the 12 transmembrane domains in AMTB is not present its plant or fungal homologues, so that an 11 transmembrane topology is predicted for the eukaryotic members of the AMT/MEP family (Thomas et al., 2000). The implication that the N-terminus of the eukaryotic AMT/MEP proteins must be on the outside of the membrane has been confirmed experimentally for the yeast MEP2 protein (Marini and Andre, 2000). Figure 6 shows a 2-D representation of the likely membrane topology of AtAMT1.1 transporter. According to this model, the majority of positively charged amino acids on the AtAMT1.1 polypeptide are on the cytoplasmic side of the membrane (Thomas et al., 2000), consistent with the ‘positive-inside’ rule (Von Heijne, 1992).

The kinetic properties of some of the Arabidopsis AMT transporters have been studied in the yeast heterologous expression system (Ninnemann et al., 1994; Gazzarrini et al., 1999; Sohlenkamp et al., 2000; Shelden et al., 2001). The ability of most AMT proteins to transport methylammonium enables (14C)methylammonium to be conveniently used to assay transport activity. The pH optimum for methylammonium uptake by yeast cells expressing AtAMT1.1 was 7, with only low rates of uptake at pH 9, consistent with NH₄⁺ being the substrate rather than NH₃ (Ninnemann et al., 1994). Transport activity was dependent on an energy source (glucose) and was sensitive to protonophores (which collapse the H+ gradient across the plasma membrane) and to inhibitors of the H+ pump (Ninnemann et al., 1994). AtAMT1.1 displayed Michaelis-Menten kinetics for methylammonium uptake, with a K_m of 65 µM. As expected, NH₄⁺ acted as a strong competitive inhibitor of methylammonium uptake (K_i of <10 µM), but AtAMT1.1 showed no affinity for K+ (Ninnemann et al., 1994). Thus AtAMT1.1 was demonstrated to be an energy-dependent high-affinity NH₄⁺ uptake system.

Subsequent studies have used the same yeast expression system to examine the kinetic properties of AtAMT1.1, AtAMT1.2, AtAMT1.3 and AtAMT2 (see Table 3 for a summary). Gazzarrini and colleagues reported that of the three AtAMT1 proteins, AtAMT1.1 had the highest affinity for NH₄⁺ (K_m 0.5µM) (Gazzarrini et al., 1999). In contrast, a more recent study (Shelden et al., 2001) found no significant difference between AtAMT1.1.and AtAMT1.2 in their affinity for NH₄⁺ and reported that AtAMT1.2 showed biphasic kinetics for methylammonium uptake, with K_m values of 36 µM and 3 µM.

AtAMT2 expressed in yeast was unable to transport methylammonium, making analysis of its biochemical properties difficult. However, it was established that yeast cells expressing AMT2 were able to use NH₄⁺ as a N source and to deplete NH₄⁺ from medium containing concentrations <100 µM, although at a slower rate than cells expressing AtAMT1.1 (Sohlenkamp et al., 2000).

Regulation of the AMT Genes

In addition to their differing biochemical properties, the AMT genes also show contrasting patterns of mRNA expression (Table 1). All four AMT genes so far analysed are expressed in roots (Ninnemann et al., 1994; Gazzarrini et al., 1999; Rawat et al., 1999; Sohlenkamp et al., 2000; Shelden et al., 2001), but only AMT1.1 and AMT2 are also expressed strongly in shoots (Gazzarrini et al., 1999; Sohlenkamp et al., 2000; Shelden et al., 2001). In roots, AtAMT1.1 and AtAMT2 are highly responsive to the NH₄⁺ supply, both being strongly repressed in the presence of high external NH₄⁺ concentrations (Rawat et al., 1997; Gazzarrini et al., 1999) (Sohlenkamp et al., 2000; Shelden et al., 2001). AtAMT1.3 is similarly regulated, but to a lesser degree, while AtAMT1.2 is constitutively expressed (Gazzarrini et al., 1999; Shelden et al., 2001).

Of the three N-responsive genes, it is AtAMT1.1 that responds to the N supply in a manner that most closely parallels the changes in HATS activity (Gazzarrini et al., 1999; Rawat et al., 1999; Shelden et al., 2001). For example, when plants were transferred from high NH₄⁺ concentrations to low, AtAMT1 mRNA levels began to increase within 1 h and continued to increase over a 24 h period concurrently with an increase in 13NH₄⁺ influx rates (Rawat et al., 1999). Similarly AtAMT1.1 mRNA levels in roots increased markedly during a 2 d period after the N supply was withdrawn, as did the 15NH₄⁺ influx rate, but there was no significant increase in AtAMT1.3 mRNA abundance during the same period (Gazzarrini et al., 1999). Nevertheless, these correlations need to be treated with caution since there are indications that post-transcriptional as well as transcriptional controls can operate in the regulation of NH₄⁺ transporter activity (see below).

There is evidence that glutamine is a key regulatory metabolite involved in the feedback regulation of AtAMT1.1 expression (Rawat et al., 1999). Strong negative correlations were found between glutamine concentrations in root extracts and both 13NH₄⁺ influx rates and AtAMT1.1 transcript levels. Furthermore, the repressive effect of NH₄⁺ on AtAMT1.1 expression could be relieved by blocking the conversion of NH₄⁺ to glutamine with methionine sulfoximine (MSX, an inhibitor of glutamine synthetase activity).

Are Post-Transcriptional Controls Involved in the Regulation of AMT Transporters?

There are several lines of evidence pointing to the possibility that regulation of the NH₄⁺ transporters is complex and operates at multiple levels. Gazzarrini and colleagues found that when Arabidopsis plants were transferred from NH4NO3 to KNO3, the NH₄⁺ influx rate increased over the following three days without any change in AtAMT1.1, AtAMT1.2 or AtAMT1.3 mRNA levels (Gazzarrini et al., 1999). Under similar conditions it was reported that the K_m for NH₄⁺ of the HATS changed from 46 µM to 7 µM (Shelden et al., 2001). Only two explanations for these observations seem possible: either one of the other AMT genes is responsible for these changes in the activity and kinetic properties of HATS (and is regulated by the N supply in a very different way from AtAMT1.1, AtAMT1.2 or AtAMT1.3), or regulation of AMT expression at some post-transcriptional level is occurring.

In experiments where MSX was used to block NH₄⁺ assimilation and NH₄⁺ consequently failed to cause a significant decline in AtAMT1.1 transcript levels, there was nevertheless a 30% decrease in 13NH₄⁺ influx rates (Rawat et al., 1999). Further evidence for post-transcriptional controls comes from experiments where the resupply of NH4NO3 to N-starved plants consistently led to a more rapid decline in NH₄⁺ influx rates than in AtAMT1.1 mRNA abundance (Rawat et al., 1997), even though AtAMT1.1 is the most highly responsive of the AMT gene family (Gazzarrini et al., 1999); (Rawat et al., 1999; Sohlenkamp et al., 2000; Shelden et al., 2001).

All of these observations are consistent with a role for the NH₄⁺ ion in the post-transcriptional regulation of the HATS (Rawat et al., 1999), perhaps by directly inhibiting AMT transporter activity or by inhibiting the synthesis or post-translational processing of the AMT proteins. A model for the feedback regulation of AtAMT1.1 by glutamine and NH₄⁺ is shown in Figure 7.

Concluding Remarks on AMT Transporter Function

The current evidence strongly suggests that one or more of the AtAMT genes encode the HATS for NH₄⁺ in Arabidopsis roots. Their differing affinities for NH₄⁺ and responsiveness to N, and the varied patterns of expression within the plant, suggest that they perform different, but perhaps overlapping, physiological functions. The high-affinity of AtAMT1.1 for NH₄⁺ and the close correlation between AtAMT1.1 mRNA levels and the HATS activity has made this member of the AMT family the strongest candidate for being a major component of the high-affinity NH₄⁺ uptake system, particularly in NO₃⁻—fed and N-deprived plants. The very high affinity of the AtAMT1.1 transporter expressed in yeast (Table 3), and its strong derepression in N-deprived roots, suggests that it accounts for the highest affinity component of the HATS (K_m of 8 µM) that has only been detected in fully NH₄⁺-deprived roots (Shelden et al., 2001).

Nevertheless, since AtAMT1.2, AtAMT1.3 and AtAMT2 are also expressed in roots, and since each has been shown to have the capacity to transport NH₄⁺ in the concentration range typical of the HATS (Table 3), it is likely that they too contribute to the HATS. The extent to which each contributes seems certain to vary with such factors as the rate and nature of the external N supply, the time of day and the N and C status of the plant. Examination of the TIGR Gene Index for Arabidopsis ESTs (Quackenbush et al., 2001) reveals that the Columbia root EST collection contains two cDNAs for AtAMT1.1, four for AtAMT1.2 and one for AtAMT1.4, which may give some indication of the relative levels of expression of the AMT genes in NH4NO₃⁻grown plants of this ecotype.

The physiological roles of the different AtAMT transporters will become clearer once we have more information about their subcellular localizations and their cellular distributions within the root and shoot. Most important will be the analysis of mutants and transgenic lines in which the expression of single and multiple AtAMT genes is modified.

Low-affinity NH₄⁺ uptake

The non-saturable kinetics of NH₄⁺ influx at external NH₄⁺ concentrations above 1 mM suggest that it is likely to be occurring by diffusion, but it is only at high external pH that diffusion of NH₃ is likely to contribute much to the low-affinity phase. In yeast, one member of the AMT/MEP family (Mep3p) has a very low affinity for NH₄⁺ (1.4–2.1 mM), so that within this one family a full range of substrate affinities is possible. The only indication so far that a member of the AtAMT family may contribute to the LATS is the report that AtAMT1.2 has a dual affinity for methylammonium when expressed in yeast, the low affinity phase having a K_m of 3 mM (Shelden et al., 2001).

Low-affinity NH₄⁺ uptake might also occur as a by-product of the activity of K+ channels or K+ transporters (Forde and Clarkson, 1999; von Wiren et al., 2001). K+ uptake systems in maize roots included both NH₄⁺-sensitive and NH₄⁺-insensitive components (Vale et al., 1987). Analysis of a mutant carrying an insertion in the gene for an inward-rectifying K+ channel (AKT1) has confirmed that a similar situation exists in Arabidopsis (Spalding et al., 1999). Furthermore, it was shown that the NH₄⁺-sensitive component corresponded to the non-AKT1 fraction of K+ transport activity (Spalding et al., 1999). One class of K+ channels that could contribute to low-affinity NH₄⁺ influx are the voltage-independent non-selective channels (VICs; (Amtmann and Sanders, 1999)). Evidence that these are permeable to NH₄⁺ has come from lipid bilayer studies on a voltage-independent K+ channel from rye roots (White, 1996) and from in situ analysis of VICs in the plasma membrane of maize root cortical cells (A. Amtmann, unpublished results). Which genes encode VICs are still unknown. Figure 4 summarises the possible pathways for high and low-affinity NH₄⁺ transport across the plasma membrane.

Ammonium sensing

In yeast, the NH₄⁺ transporter Mep2p acts as a sensor for environmental NH₄⁺ in regulating the initiation of the process of pseudohyphal differentiation (Lorenz and Heitman, 1998). Thus Mep2p belongs to a class of membrane transporters in yeast that also serve as sensors for metabolites; other examples include the Snf3p and Rgt2p glucose transporters (Ozcan et al., 1996) and the Ssy1p and Ptr3p amino acid transporters (Klasson et al., 1999), which each sense the extracellular levels of their respective substrates. Often transporters that function as nutrient sensors have extended N- or C-terminal domains that may serve as signalling domains. None of the AMT sequences possess any additional domains that might suggest a role other than in solute transport, but since this also true of Mep2p it is still a possibility worthy of investigation that one or more of the AMT permeases function as NH₄⁺ sensors.

Nitrate Stimulation of Lateral Root Growth

A series of classical studies in the 1970′s examined the effects of NO₃⁻ and NH₄⁺ on root development in barley (Hordeum vulgare) (Drew et al., 1973; Drew, 1975; Drew and Saker, 1975). Using sand cultures or flowing nutrient solutions, Drew and colleagues found that locally concentrated supplies of NO₃⁻, NH₄⁺ or phosphate stimulated root branching specifically within the nutrient-rich zones. The response consisted of a stimulation of both lateral root initiation and lateral root growth in the zone of treatment. In contrast, growth of the seminal roots was little affected by the external nutrient supply (Drew et al., 1973), showing that different types of root meristem within one root system can have very different sensitivities to nutrients.

The development of a technique for applying nutrients in a localised fashion to Arabidopsis seedlings growing on vertical agar plates (Figure 8) allowed this phenomenon to be investigated in a species more amenable to molecular genetics (Zhang and Forde, 1998). Initial studies showed that localised NO₃⁻ treatments did indeed stimulate lateral root elongation (2- to 3-fold), specifically within the NO₃⁻—rich zone (Figure 9). The stimulation occurred even if the NO₃⁻ treatments were delayed until after the lateral roots had emerged (Zhang and Forde, 2000), showing that the NO₃⁻ does not need to be present during initiation or maturation of lateral root primordia for it to be effective, as has also been shown in wheat (Hackett, 1972). As in barley and other species, primary root growth in Arabidopsis is remarkably insensitive to changes in the NO₃⁻ supply (Zhang and Forde, 1998).

Neither the number of lateral roots initiated (Zhang and Forde, 1998), nor the rate of progression through the early stages of lateral root development (Zhang and Forde, 2000), are modified by localised NO₃⁻ treatments. The absence of any significant effect on lateral root initiation differs from the situation in barley and wheat, where the localised response to NO₃⁻ consisted of both an increase in elongation rates and an increase in numbers of laterals (Hackett, 1972; Drew et al., 1973). In Arabidopsis it was shown that even relatively low concentrations of NO₃⁻ (50–100 µM) in the NO₃—enriched zone were sufficient to elicit the response (Zhang et al., 1999). The absence of any significant increase in the size of mature cells in the NO₃⁻—stimulated roots established that the increased growth rates were due to increased meristematic activity in the lateral root tip (Zhang et al., 1999).

When localised supplies of NH₄⁺ or glutamine were substituted for NO₃⁻, no stimulation of lateral root growth was observed (Zhang et al., 1999), indicating that the roots were responding to the NO₃⁻ ion itself rather than to NO₃⁻ as a source of N. Further evidence for this was obtained using a nia1nia2 mutant which is deficient in NR and hence unable to use NO₃⁻ as an effective N source (Wilkinson and Crawford, 1993); this mutant showed a wild-type response to a localised NO₃⁻ treatment (Zhang and Forde, 1998). Thus, just as NO₃⁻ is a signal for the induction of the nitrate assimilatory pathway (see above), it is also a signal for increased root proliferation. It seems that individual lateral root tips have a mechanism for monitoring the external NO₃⁻ concentration and for modifying their meristematic activity accordingly (Zhang and Forde, 1998; Zhang et al., 1999).

Genes Involved in Nitrate Stimulation of Lateral Root Growth

A possible component of the signal transduction pathway linking external NO₃⁻ to changes in meristematic activity has been identified during a screen for NO₃⁻ regulated genes in Arabidopsis roots (Zhang and Forde, 1998). Amongst the novel NO₃⁻—inducible sequences identified was ANR1, a member of the MADS box family of transcription factors (Zhang and Forde, 1998). There are at least 82 MADS box genes in the Arabidopsis genome (Riechmann et al., 2000) and most of those that have so far been functionally characterised are involved in controlling floral organogenesis or other processes related to the reproductive phase of growth (Ng and Yanofsky, 2001). ANR1 belongs to a group of nine MADS box genes so far known to be expressed in Arabidopsis roots (Alvarez-Buylla et al., 2000), but most of these are of unknown function. When transgenic lines were generated in which ANR1 expression was down-regulated (by antisense or co-suppression), their lateral roots were found to have lost the ability to respond to localised supplies of NO₃⁻ (Figure 9). It was proposed that ANR1 is a component of the sensory pathway linking NO₃⁻ to the stimulation of lateral root growth (Zhang and Forde, 1998).

Auxin is well known as an important regulator of root growth (Leyser, 1997), and experiments using auxin-response mutants of Arabidopsis have provided evidence for an overlap between NO₃⁻ signalling and auxin signalling in the control of lateral root elongation (Zhang et al., 1999). Three auxin-response mutants (aux1, axr2 and axr4) were tested for their sensitivity to localised supplies of NO₃⁻: While aux1 and axr2 showed responses that were not significantly different from the wild-type, the axr4 mutant showed a phenotype similar to the ANR1 down-regulated lines (Zhang et al., 1999). Although the AXR4 gene has not been identified at the molecular level, its location on chromosome 1 (Hobbie and Estelle, 1995) establishes it as distinct from ANR1 (which is on chromosome 2). Unusual among auxin-response mutants, the phenotype of axr4 mutants is restricted to roots and does not involve cross-resistance to other plant hormones such as cytokinin or ethylene (Hobbie and Estelle, 1995). Apart from reduced gravitropism, the root phenotype of the axr4-2 mutant is remarkably like the wild-type, making its altered sensitivity to NO₃⁻ stimulation of lateral root elongation of particular interest. A model for the NO₃⁻ signalling pathway leading to increased meristematic activity in the lateral root tip is shown in Figure 10.

Nitrate as an Inhibitor of Early Lateral Root Development

Nitrate has also been found to have an inhibitory effect on lateral root development (Scheible et al., 1997c; Zhang and Forde, 1998; Stitt, 1999; Zhang et al., 1999). In Arabidopsis this consists of a delayed outgrowth of the lateral root primordium when roots are exposed to NO₃⁻ concentrations above 1 mM (Zhang et al., 1999). At 50 mM, the highest NO₃⁻ concentration tested, two-week-old seedlings developed primary roots of the same length as at 1 mM, and had a similar number of lateral root primordia, but most or all of the laterals were less than 0.5 mm in length. This inhibitory effect of NO₃⁻ appears to be very different in nature from its stimulatory effect (Zhang et al., 1999; Zhang and Forde, 2000). Firstly, while the stimulatory effect acts on elongation of mature laterals, the inhibitory effect acts specifically on immature LRs during a discrete phase around the time of their emergence from the primary root. Secondly, the high external NO₃⁻ concentration does not act directly on the developing lateral; if only a portion of the primary root was exposed to 50 mM NO₃⁻, the inhibitory effect was much diminished and what inhibitory effect remained was distributed along the root (Zhang et al., 1999). Thus the effect seems to depend on the NO₃⁻ absorbed by the plant rather than the external NO₃⁻ concentration per se.

Using the NR-deficient nia1nia2 mutant it was shown that the assimilation of NO₃⁻ was not required for the inhibitory effect, and furthermore that lateral root development in the mutant was rather more sensitive to NO₃⁻ inhibition than the wild-type (Zhang et al., 1999). Nitrate reductase-deficient mutants growing on NO₃⁻ are known to accumulate high levels of tissue NO₃⁻, particularly in the leaves e.g. Warner and Huffaker, 1989), so this observation is consistent with the notion that NO₃⁻ levels inside the plant are responsible for generating the inhibitory effect on lateral root outgrowth. A similar phenomenon has been reported in tobacco, where NR deficient lines growing on NO₃⁻ display a marked inhibition of root growth (Scheible et al., 1997b; Scheible et al., 1997c) due to a reduction in lateral root frequency (Stitt, 1999). Here the evidence from metabolite analysis and split root experiments has lead to the conclusion that the inhibition of root growth is due to a signal from the shoot triggered by NO₃⁻ accumulation in the shoot (Scheible et al., 1997c). In Arabidopsis (but not in tobacco), an increase in the sucrose concentration in the medium led to a partial alleviation of the inhibitory effect of NO₃⁻ on root branching (Scheible et al., 1997c; Zhang et al., 1999). This would suggest that either the balance between the carbon and nitrogen status of the plant is being monitored, or that sucrose itself has a signalling role that acts antagonistically with the NO₃⁻ signal (Smeekens, 2000).

Why outgrowth of the lateral root primordium should be particularly sensitive to inhibition by NO₃⁻ is unclear. From other work it is known that the stage just after its emergence is a critical step in the development of a lateral root (Cheng et al., 1995; Malamy and Benfey, 1997a; Malamy and Benfey, 1997b). Although the lateral root meristem appears to be fully formed prior to emergence, emergence itself takes place through cell expansion rather than cell division (Malamy and Benfey, 1997b). It is only after emergence that activation of the lateral root meristem occurs and growth of the mature lateral root begins (Cheng et al., 1995; Malamy and Benfey, 1997b). Thus there may be a developmental process between differentiation of the lateral root meristem and its activation that is particularly sensitive to high NO₃⁻ concentrations. An alternative explanation could lie in the timing of the establishment of symplastic connections between the phloem of the primary root and the lateral root primordium. Since a shoot-derived signal may be involved, these connections could be necessary for transmission of the signal to the developing primordium. A study using fluorescent dyes and confocal microscopy has elegantly demonstrated for Arabidopsis that early in its development the lateral root is symplastically isolated and that the earliest functional symplastic connections with the lateral root primordium occur around the time of emergence from the cortex (Oparka et al., 1995).

The finding that NO₃⁻ can either inhibit or stimulate lateral root production, depending on its external distribution and its accumulation within plant tissues, has led to a ‘dual pathway’ model for NO₃⁻ regulation of root branching in Arabidopsis (Zhang et al., 1999). This model, illustrated in Figure 11, depicts one signalling pathway (the localised stimulatory pathway) which responds to the external NO₃⁻ concentration at the lateral root tip and regulates increase meristematic activity specifically in that root tip, and a second signalling pathway (the systemic inhibitory pathway) which responds to the internal NO₃⁻ status of the plant and regulates progress through the latter stages of lateral root maturation. These two antagonistic pathways for regulation of root branching can be seen as providing a mechanism by which the plant can modify its root architecture in a manner that integrates information about the spatial distribution of NO₃⁻ in the soil and the plant's current N status (Zhang et al., 1999). Thus the plant could modulate the intensity of its proliferative response to a localised NO₃⁻ supply according to its demand for N. This would allow the allocation of resources within the plant to be optimised by ensuring that the strongest response to a localised NO₃⁻ supply occurred when the plant was N-depleted (Zhang et al., 1999). Increased lateral root proliferation is only one component of a plant's response to localised NO₃⁻ supplies; a much faster response is seen at the metabolic level, where NO₃⁻ influx by roots located within a NO₃⁻—rich patch is rapidly induced (Laine et al., 1995; Laine et al., 1998). The up-regulation of the NO₃⁻ influx system can be seen as a short-term adaptive response to localised NO₃⁻, with stimulation of lateral root development being the longer-term component of the response. It is known that the NO₃⁻ influx system is regulated by signals originating in the shoot (Laine et al., 1995), and it remains to be seen whether the same signals are involved in regulating lateral root development.

Ammonium Inhibition of Root Growth

Ammonium, when supplied to plants as the sole N source, often inhibits plant growth, particularly in unbuffered media when acidification of the rhizosphere becomes a problem (von Wiren et al., 2001). However, a novel aspect of NH₄⁺ inhibition of root growth has been reported in Arabidopsis (Cao et al., 1993). In these studies it was found that if K+ was omitted from the medium, and NH₄⁺ concentrations were high (>3 mM) seedling root growth was dramatically inhibited, even when NO₃⁻ was present. This inhibition could be reversed if low levels of K+, Rb+ or Cs+ ions were added, and the protective effect of K+ was shown not to be due to an inhibition of NH₄⁺ uptake. The further finding that three auxin-response mutants (aux1, axr1 and axr2) were resistant to NH₄⁺ inhibition of root growth led to the suggestion that this unusual response to NH₄⁺ was mediated by effects on hormone metabolism or signalling. Since it is unlikely that an Arabidopsis plant will often be exposed in nature to the required combination of high NH₄⁺ and very low K+ concentrations, it is unclear what the physiological significance of this intriguing phenomenon may be; however, it is an example in Arabidopsis showing the role of auxin in nutrient effects on root growth.

Nitrogen Regulation of Shoot Growth and Development

It has been established for Arabidopsis that a higher rate of N supply leads to an increased allocation of resources to shoot growth (Schulze et al., 1994), leading to increased shoot:root ratios, which is the morphological response commonly seen in many other plant species (Ericsson, 1995). Analysis of metabolite pools showed that there was a striking correlation between the concentrations of NO₃⁻ and organic N in rosette leaves and shoot growth rates (Schulze et al., 1994). In tobacco, transgenic lines with low NR activity have helped to demonstrate the role of the shoot NO₃⁻ pool as a signal in regulating shoot-root allocation (Scheible et al., 1997c).

Other studies have highlighted the importance of the short-term N supply and root-to-shoot signalling in the regulation of leaf growth (McDonald and Davies, 1996). Recent studies with tobacco found that the rate of cell division and cell elongation in leaves fell markedly within 24 h of transferring plants from NO₃⁻ to NH₄⁺ (Walch-Liu et al., 2000). This effect was not due to limitation of carbohydrates or N, nor to a lack of osmotica or water. A possible role for NO₃⁻ as a signal was inferred from evidence that it was absence of NO₃⁻ rather than the presence of NH₄⁺ that was responsible for the reduction in leaf growth (Walch-Liu et al., 2000). A decline in the zeatin + zeatin riboside fraction in xylem exudates which paralleled that in leaf expansion rates led to the suggestion that NO₃⁻ may be involved in a signal transduction chain that regulates leaf morphogenesis by modulating the flux of cytokinin from root to shoot (Walch-Liu et al., 2000).

Although leaves of NO₃⁻—grown Arabidopsis plants are larger than those of NH₄⁺-grown plants (e.g. (Oostinder-Braaksma and Feenstra, 1973)), the possible short-term modulation of leaf expansion by the NO₃⁻ supply has not been studied in Arabidopsis. However it is interesting to note that some members of the Arabidopsis family of response regulators (ARRs) have been shown to be induced in leaves by either the direct application of cytokinins or the re-supply of NO₃⁻ to roots of N-starved plants (Taniguchi et al., 1998). It has been proposed that the ARR genes are part of a cytokinin-mediated signalling pathway which enables gene expression in leaves to be modulated by the inorganic N supply to the roots (Sakakibara et al., 2000). The finding that one of the cytokinin-regulated genes (ARR5) is expressed in shoot apical meristems (D'Agostino et al., 2000), would be consistent with a possible role in regulating leaf expansion in response to the NO₃⁻ supply. A tentative model for such a NO₃⁻—cytokinin signalling pathway is depicted in Figure 12.

In many species, increased N fertilisation is reported to delay flowering (Bernier et al., 1993). In one study, flowering time in Arabidopsis was found to be only slightly delayed by a high rate of N supply (6 mM compared to 0.1 mM NH4NO3) (Schulze et al., 1994). In this study, seed set was much more sensitive to the N supply than flowering time, with the number of seeds per plant being over 10-fold greater under the high N regime. Importantly, by switching plants from high N to low N and vice versa at the end of the vegetative phase of growth, it was shown that it was the N supply during the reproductive phase that was critical in determining seed numbers (Schulze et al., 1994). It is not yet known which are the key N pools responsible for regulating leaf growth and seed set in Arabidopsis.