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First published online August 19, 2004; 10.1105/tpc.104.024380 © 2004 American Society of Plant Biologists
Transcript Profiling in the chl1-5 Mutant of Arabidopsis Reveals a Role of the Nitrate Transporter NRT1.1 in the Regulation of Another Nitrate Transporter, NRT2.1
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ABSTRACT |
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 TOP ABSTRACT INTRODUCTIONRESULTSDISCUSSIONMETHODSREFERENCES |
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INTRODUCTION |
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TOPABSTRACTINTRODUCTIONRESULTSDISCUSSIONMETHODSREFERENCES |
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) and AMT1 and AMT2 families for NH4+ transporters (von Wirén et al., 2000). Concerning more specifically NO3– transport in Arabidopsis thaliana, the NRT2 family includes seven genes (Orsel et al., 2002), but the NRT1 family is more difficult to define precisely. It has been restricted to four genes in previous studies (Okamoto et al., 2003), but these genes belong to the large PTR family of transporters, with 51 members. To date, nothing excludes the possibility that NO3– transporters are also encoded by other PTR genes than the four NRT1 initially investigated. Of these 58 putative transporters (seven NRT2 and 51 PTR), only three (NRT1.1, NRT1.2, and NRT2.1) have been functionally characterized in planta and shown to ensure part of the NO3– uptake from the external medium. NRT1.1 (also called CHL1) was the first NO3– transporter identified in plants (Tsay et al., 1993) and was initially believed to be a NO3– inducible low-affinity transporter (Tsay et al., 1993; Huang et al., 1996; Touraine and Glass, 1997). In the same family as NRT1.1, NRT1.2 was also characterized as a low-affinity NO3– transporter, but with a constitutive expression, not dependent on the presence of NO3– (Huang et al., 1999). Thus, both NRT1.1 and NRT1.2 were considered to belong to the NO3– LATS in A. thaliana (Crawford and Glass, 1998). These two transporters do not fulfill similar functions because NRT1.1 mutants appear to be strongly defective in LATS activity only when plants are supplied with a mixed NO3– + NH4+ N source (Touraine and Glass, 1997; Crawford and Glass, 1998), whereas antisense lines of NRT1.2 also display a markedly reduced LATS activity on NO3– as sole N source (Huang et al., 1999). On the other hand, the NRT2.1 gene was shown to encode a major component of the NO3– HATS (Lejay et al., 1999; Zhuo et al., 1999; Cerezo et al., 2001; Filleur et al., 2001). It appears to play a crucial role in the control of the high-affinity NO3– uptake by the plant because its expression is regulated as the NO3– HATS is. For example, it is inducible by NO3– itself (Filleur and Daniel Vedele, 1999; Zhuo et al., 1999; Nazoa et al., 2003), repressed by reduced N metabolites (Lejay et al., 1999; Zhuo et al., 1999), and stimulated by photosynthates (Lejay et al., 1999, 2003). Although NRT1.1, NRT1.2, and NRT2.1 do not mediate all NO3– transport steps occurring in the plant, these three transporters have a central importance for NO3– acquisition in A. thaliana. Hence, in addition to the study of some of the other 55 putative transporters (Orsel et al., 2002; Okamoto et al., 2003), further analysis of the overall function of NRT1.1, NRT1.2, and NRT2.1 is still needed. Concerning NRT1.1, the picture becomes more and more complex with recent reports indicating that the role of this transporter is far from being limited to low-affinity NO3– uptake in roots. First, NRT1.1 is now considered as a dual affinity transporter, belonging to both HATS and LATS (Wang et al., 1998; Liu et al., 1999). Phosphorylation of NRT1.1, triggered by limited external NO3– availability, is responsible for the shift from low to high affinity, thus adapting the functional properties of the transporter to the resource level in the root environment (Liu and Tsay, 2003). Second, NRT1.1 is strongly expressed in nascent organs of both root and shoot (root tip, emerging lateral roots, and nascent leaves) and plays a crucial role in the early phases of development of these young organs (Guo et al., 2001). In particular, several NRT1.1 mutants display altered root architecture in some conditions, with a reduced growth of both primary and secondary roots, sometimes even in the absence of added NO3– in the external medium. This suggests an alternative function for NRT1.1, independent of NO3– transport (Guo et al., 2001). Finally, it has been reported recently that the mutation of NRT1.1 also leads to a lower sensitivity to drought, related to reduced stomatal opening because of impaired NO3– transport in stomata guard cells (Guo et al., 2003). Clearly, the view that NRT1.1 behaves only as a transporter in charge of the NO3– uptake from the external medium is an oversimplification. This protein appears to fulfill a multiplicity of physiological functions, which begin to be unravelled more than 30 years after the identification of the first NRT1.1 mutant (Oostindiër-Braaksma and Feenstra, 1973).
Most of the novel and important findings mentioned above concerning NRT1.1 rely on physiological or morphological analyses of mutants. Very few molecular data are associated with these reports, thus resulting in a lack of understanding of the gene networks functionally associated with NRT1.1 in the control of N acquisition, root and shoot development, and water use in the plant. To obtain further insight on this point, we performed large-scale transcript profiling in roots of both the chl1-5 mutant of NRT1.1 (Tsay et al., 1993) and the related Columbia (Col) wild type. Our transcriptomic approach was based on the serial analysis of gene expression (SAGE) methodology (Velculescu et al., 1995), which involves the generation of a short specific tag (14 bp) for each mRNA in a sample. The sequencing of a large number of SAGE tags in a sample library allows a high-throughput analysis of the frequencies of these tags, which are representative of the relative amounts of the corresponding mRNAs. Thus, the comparison of the tag sequences and copy numbers obtained from two different libraries allows the identification of the genes differentially expressed between the two original samples. SAGE has been mostly employed in cancer research (Boon et al., 2002) but is now increasingly used in plants (Lorenz and Dean, 2002; Matsumura et al., 2003), especially in A. thaliana (Jung et al., 2003; Lee and Lee, 2003), in which the full genome sequence provides a unique tool for identifying the genes corresponding to the tags found experimentally (Fizames et al., 2004).
In addition to the finding that many genes show a markedly altered level of expression in the roots of the chl1-5 mutant as compared with the Col wild type, we report here the observation that NRT2.1 expression is markedly deregulated in the mutant, a response that could not be explained by the known regulation affecting this gene. This suggests either the occurrence of a yet unknown signaling for control of NRT2.1 expression or a role of NRT1.1 in the regulation of other NO3– transporters at the gene expression level.
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RESULTS |
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TOPABSTRACTINTRODUCTIONRESULTSDISCUSSIONMETHODSREFERENCES |
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; Crawford and Glass, 1998), the two SAGE libraries were generated from roots of Col-0 and chl1-5 plants grown hydroponically on 1 mM NH4NO3. These libraries were sequenced up to 31,354 and 28,451 tags for Col-0 and chl1-5 roots, respectively (the Col-0 library has already been reported under the name of the NH4NO3 library in Fizames et al., 2004). The 59,805 total tags correspond to 25,230 different sequences, among which 7583 are represented by tags found at least twice and up to 228 times. Because of rare but unavoidable sequencing and PCR errors, the use of single tags is not totally safe for gene identification. Thus, we restricted our analysis to the 7583 different tags found at least twice in the two combined libraries. The identification of the genes represented in our SAGE transcriptomes was performed by matching the list of the 7583 experimental tags to that of the virtual ones obtained by extracting the predicted SAGE tag sequence from each gene annotated in the whole Arabidopsis genome (Fizames et al., 2004). Among the 7583 different experimental tags, 1972 had no gene match in the database of virtual SAGE tags, 885 matched several genes and were thus not specific, and 4726 matched one single gene. The whole set of data on the 5611 tags matching one or several genes can be viewed online (http://genoplante-info.infobiogen.fr; see Supplemental Table 1 online). The statistical analysis of the comparison between Col and chl1-5 libraries (Figure 1) resulted in the identification of 419 tags with different copy numbers in the two libraries at P < 0.01 (1194 tags at P < 0.05). Among these, 296 tags could be attributed to one single gene at P < 0.01 (797 tags at P < 0.05). The 296 differentially expressed genes (P < 0.01) reveal a large variety of functions affected in the chl1-5 mutant (http://genoplante-info.infobiogen.fr; see Supplemental Table 2 online) but also include genes directly related either to N nutrition or ion transport (Table 1). A few genes encoding enzymes of N metabolism have a strongly altered expression. This is the case of two isoforms of glutamate dehydrogenase (GDH1 and GDH2), which are markedly underexpressed in roots of chl1-5 compared with Col. Several transporter or channel genes also show changes in expression between the two genotypes. Of particular interest are those related to NO3– or amino acid transport, such as NRT2.1, At1g32450 (a member of the large PTR multigene family including NRT1.1), At4g38250, and AAP6, which are all strongly overexpressed in chl1-5. On the other hand, genes encoding aquaporins (PIP2;2, PIP1;2, PIP2;1, and PIP1;1) and metal (IRT1 and NRAMP1), SO42– (SULTR1;2), or K+ (SKOR) transporters/channels are repressed in the mutant.
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), NIA1 and NIA2 (encoding the two nitrate reductase [NR] isoforms present in A. thaliana; Wilkinson and Crawford, 1993), GS2 (encoding the chloroplastic isoform of Gln synthetase; Peterman and Goodman, 1991), AMT1.1 (encoding a high-affinity NH4+ transporter; Ninnemann et al., 1994), SKOR (encoding a K+ channel implicated in xylem loading; Gaymard et al., 1998), and IRT1 (encoding an iron transporter; Vert et al., 2002). The absence of the NRT1.1 SAGE tag in the chl1-5 library could not be verified because this tag is not specific and also matches nine other genes, but as expected, NRT1.1 transcript was not detected in the chl1-5 mutant (data not shown). NIA1, NIA2, GS2, and AMT1.1 were not found in the list of differentially expressed genes and did not show any significant difference in their transcript accumulation between Col and chl1-5 roots (Figure 2). The slightly higher mRNA levels apparent in Figure 2 for NIA1 and NIA2 in the mutant are not representative. In four independent experiments, the chl1-5/Col ratio was measured at 0.81 ± 0.25 and 1.37 ± 0.38 for transcript accumulation of NIA1 and NIA2, respectively. At the opposite, NRT2.1 was found by SAGE to be significantly overexpressed in chl1-5 roots as compared with Col roots (Table 1). This marked difference in transcript accumulation was confirmed by RNA gel blot analysis (Figure 2), with approximately the same ratio of six between Col and chl1-5 (mean value for this ratio in five independent experiments: 6.51 ± 1.76). For both SKOR and IRT1, the SAGE data suggested a lower expression level in roots of chl1-5 than in those of Col, which was also verified by RNA gel blot analysis (Figure 2).
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; Zhuo et al., 1999; Cerezo et al., 2001) (i.e., under the conditions of our study). Thus, the totally unexpected observation of a high NRT2.1 expression level in chl1-5 in the presence of NH4+ prompted us to focus further investigation on this intriguing point.
Numerous reports have shown a strong correlation between NRT2.1 transcript accumulation in the roots and the activity of the HATS for NO3– (Lejay et al., 1999; Zhuo et al., 1999; Cerezo et al., 2001; Gansel et al., 2001; Okamoto et al., 2003). Thus, we investigated if NRT2.1 overexpression in the chl1-5 mutant also had functional consequences on NO3– uptake rate by this mutant. To do so, the kinetics of 15NO3– influx as a function of external 15NO3– concentration was determined in both Col and chl1-5 plants grown for 6 weeks on 1 mM NH4NO3 (Figure 3A). In the low NO3– concentration range (10 to 500 µM), 15NO3– influx in chl1-5 was higher than in Col, whereas in the high concentration range (0.5 to 5 mM), the reverse was observed with 15NO3– influx in chl1-5 roots lower than in Col. The stimulation of the HATS activity in chl1-5 as compared with Col was most pronounced at 25 to 50 µM external 15NO3– (approximately fourfold increase; Figure 3B). This is the exact range of concentration where NRT2.1 was shown to participate predominantly to root NO3– uptake (Cerezo et al., 2001), indicating that the upregulation of the HATS in chl1-5 plants was most probably attributable to the overexpression of NRT2.1 as compared with Col.
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; Touraine and Glass, 1997; Wang et al., 1998; Liu et al., 1999; Guo et al., 2001). To determine whether other genes than NRT1.1 are also absent in chl1-5, we used a PCR approach to map the deletion. Successive PCR amplifications were performed on Col and chl1-5 genomic DNA using oligonucleotides designed from the sequence of chromosome 1 (GenBank accession number NC_003070). This revealed that the chl1-5 deletion corresponds to an 18.31-kb DNA fragment, beginning in the last NRT1.1 intron and ending after the At1g12090 gene (Figure 4). These data show that not only NRT1.1 but also two other genes, At1g12090 and At1g12100, are affected in the chl1-5 mutant. These two genes are highly homologous. No EST was found for the At1g12100 gene in all available databases, and its SAGE tag was absent from both Col and chl1-5 libraries. The At1g12090 gene seems to be significantly expressed because many ESTs were found, with some of them corresponding to a root specific library. The SAGE tag of At1g12090 was recorded four times in the Col library, but as expected, was not found in the chl1-5 library.
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Regulation of NRT2.1 Expression by N Status of the Plant Is Altered in NRT1.1 Mutants
The overexpression of NRT2.1 observed in NH4NO3-grown chl1-5 plants was also found in three other NRT1.1 mutants (Figure 5): chl1-10, chl1-11, and the original chl1-1 mutant (formerly called B1; Doddema and Telkamp, 1979). This demonstrates that upregulation of NRT2.1 expression is specifically attributable to the NRT1.1 mutation. Moreover, the comparison of wild-type and mutant plants either grown on NO3– or NH4NO3 showed that NRT2.1 expression was strongly repressed by NH4+ in wild-type plants, but surprisingly not in chl1-1, chl1-10, and chl1-11 plants (Figure 5).
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; Zhuo et al., 1999; Cerezo et al., 2001), upregulation in response to N starvation (Filleur and Daniel-Vedele, 1999; Lejay et al., 1999), induction by NO3– (Filleur and Daniel-Vedele, 1999), and diurnal changes (Lejay et al., 1999).
As previously described (Lejay et al., 1999; Zhuo et al., 1999; Cerezo et al., 2001), NRT2.1 transcript level was rapidly and markedly lowered in wild-type plants by exogenous NH4+ supply or high external NO3– concentration (Figure 6). This downregulation of NRT2.1 expression was absent or much less pronounced in chl1-5 plants (Figure 6). Addition of 5 mM Gln (a strong repressor of NRT2.1 expression) to the 1 mM NO3– medium resulted after 6 h in a nearly 90% decrease of NRT2.1 transcript level in Col roots, whereas this decrease was only of 50% in chl1-5 roots (data not shown). Not only the expression of NRT2.1, but also the activity of the HATS for NO3– was resistant to the repression exerted by a reduced N source in chl1-5. Root 15NO3– influx measured at 0.2 mM external concentration was lowered by 
50% after NH4+ supply in Col plants but was unaffected in chl1-5 plants (Figure 7). Another clear example of altered regulation of NRT2.1 expression in chl1-5 plants relates to the response to N starvation (Figure 8). In Col roots, NRT2.1 transcript level increased 24 and 48 h after transfer of the plants to N-free solution and decreased again thereafter. This transient upregulation has been attributed to the opposite effects of two different regulatory mechanisms (Lejay et al., 1999): relief from repression by N metabolites (initially predominant), on the one hand, and shortage of induction by NO3– after several days without NO3– supply (predominant after 2 d), on the other hand. In chl1-5 roots, the initial increase in NRT2.1 expression after removal of the N source was absent, and only the decay of NRT2.1 transcript level because of deinduction was observed (Figure 8). Most importantly, this altered response to N starvation is not found for all genes regulated by N status because AMT1.1, encoding an N starvation induced NH4+ transporter (Gazzarrini et al., 1999; Rawat et al., 1999), displayed a similar upregulation after transfer of the plants to N-free solution in both Col and chl1-5 roots (Figure 8). The two other main regulations identified for the control of NRT2.1 expression, namely induction by NO3– (Filleur and Daniel-Vedele, 1999) and diurnal changes (Lejay et al., 1999, 2003), are not affected in the chl1-5 mutant as compared with Col (Figure 9).
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DISCUSSION |
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TOPABSTRACTINTRODUCTIONRESULTSDISCUSSIONMETHODSREFERENCES |
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; Touraine and Glass, 1997; Wang et al., 1998; Liu et al., 1999; Guo et al., 2001, 2003), caution is needed in associating all these changes in gene expression to the mutation of NRT1.1. First, transcript profiling was performed in only one series of experiments because of the cost of SAGE. Second, two other genes (At1g12090 and At1g12100) were also found to be deleted in chl1-5. Of these two genes, only At1g12090 seems to be expressed in the roots. Its function is unclear because it encodes a protein sharing similarities with pEARLI, an extensin, a protease inhibitor, and AIR1, this latter being possibly involved in the auxin-mediated initiation of lateral roots (Neuteboom et al., 1999). Further work is thus needed, using other chl1 mutants (Figure 5), to determine the individual genes whose expression is specifically altered by NRT1.1 mutation. Nevertheless, some of the molecular responses observed in chl1-5 are correlated with physiological modifications reported in the ch11-1 mutant, suggesting that they result from NRT1.1 deletion. For instance, the fact that various metal/K+/SO42– transporter/channel genes were found to be downregulated in chl1-5 (Table 1) is consistent with the observation that the chl1-1 mutant is not only altered in NO3– transport but also in the uptake of several other ions (Scholten and Feenstra, 1986).
Concerning NO3– uptake, our observation of a lowered LATS activity in the chl1-5 mutant compared with Col when the plants are supplied with NH4NO3 as an N source (Figure 3) is in very good agreement with previous reports on this mutant (Huang et al., 1996; Touraine and Glass, 1997). However, we found this alteration of the LATS compensated for by a much higher HATS activity in chl1-5 than in Col (Figure 3). These results contrast with previous observations that chl1-5 and other chl1 alleles are defective in both HATS and LATS for NO3– (Wang et al., 1998; Liu et al., 1999). The reasons for this discrepancy between our results and those of Wang et al. (1998) and Liu et al. (1999) are unclear. However, impaired NO3– HATS activity in NRT1.1 mutants has always been reported in much younger plants (5 to 12 d old) than those used in our study (6 weeks old). Also, many other specific conditions (in particular carbon sources and NO3– and NH4+ concentrations) were different between our experiments and those of Wang et al. (1998) and Liu et al. (1999) and may explain these contrasting conclusions.
Although a putative NO3– transporter gene (At1g32450), uncharacterized to date, is also upregulated in chl1-5, we hypothesize that the stimulation of the NO3– HATS in the mutant is because of the overexpression of NRT2.1. This gene is believed to play a key role in the N acquisition by the roots. It encodes a major component of the HATS for NO3– in A. thaliana, and its expression is strongly regulated according to N/C status of the plant (Filleur and Daniel-Vedele, 1999; Lejay et al., 1999; Zhuo et al., 1999; Cerezo et al., 2001; Filleur et al., 2001; Gansel et al., 2001; Lejay et al., 2003). At least three major mechanisms have been proposed to explain the changes in NRT2.1 transcript accumulation in the root: induction by NO3– (Filleur and Daniel-Vedele, 1999; Zhuo et al., 1999; Nazoa et al., 2003), induction by light and sugars (Lejay et al., 1999, 2003), and feedback repression by N metabolites (Lejay et al., 1999; Zhuo et al., 1999; Gansel et al., 2001; Cerezo et al., 2001). Downregulation of NRT2.1 by N metabolites is postulated to involve products of NO3– assimilation, and more particularly NH4+ and Gln, as negative effectors of the expression of the gene (Lejay et al., 1999; Zhuo et al., 1999; Nazoa et al., 2003). This is expected to ensure the tuning of the HATS activity to the N demand of the whole plant. Accordingly, the deletion of NRT2.1 (together with part of NRT2.2) in the atnrt2 mutant results in both a lowered activity of the HATS and in the loss of the regulation of this uptake system by the N status of the plant (Cerezo et al., 2001; Filleur et al., 2001).
We show here that the regulation of NRT2.1 transcript accumulation by N status of the plant is altered in the chl1-5 mutant (Figures 2, 6, and 8), as well as in other NRT1.1 mutants (Figures 5 and 13). The first hypothesis that can be considered for explaining the increase in NRT2.1 transcript level in the NRT1.1 mutants is that these mutants suffer from N deficiency even when supplied with mixed N sources such as NH4NO3. However, several lines of evidence do not support this hypothesis. First, chl1-5 plants are not deficient for NH4+ uptake (Figure 10), and neither the total N influx in roots (NO3– plus NH4+; Figures 3 and 10), nor the total N content of both roots and shoots (data not shown) markedly differ between chl1-5 and Col plants when grown on 1 mM NH4NO3. Second, the observation that the accumulation of free Gln in roots is higher in chl1-5 than in Col (Figure 11) also clearly indicates that chl1-5 plants are N sufficient. Third, the activity of the NH4+ HATS in chl1-5 plants is not derepressed as expected if these plants were N deficient. Root 15NH4+ influx, measured at 0.2 mM external 15NH4+, is low (50 µmol h–1 g–1 root dry weight) and similar in both chl1-5 and Col (Figure 10). In N-limited plants, root 15NH4+ influx is generally recorded at much higher values (up to 500 µmol h–1 g–1 root dry weight; Gazzarrini et al., 1999; Rawat et al., 1999). Accordingly, the expression of the N starvation–inducible NH4+ transporter gene AMT1.1 (Gazzarrini et al., 1999; Rawat et al., 1999) is low in the chl1-5 mutant on 1 mM NH4NO3 and not different than in Col (Figure 8). Thus, the overexpression of NRT2.1 in chl1-5 cannot be explained by general N deficiency. This suggests that normal regulation of NRT2.1 expression by N status of the plant is markedly altered in chl1-5 plants. Indeed, submitting the plants to much more repressive conditions (transfer to 10 mM NO3–, 10 mM NH4NO3, or 5 mM Gln) did not result in a strong repression of NRT2.1 expression in chl1-5, whereas these treatments almost completely abolished it in Col (Figure 6). Reciprocally, transfer of the plants to N-deprived solution failed to derepress this gene in chl1-5, whereas the usual transient upregulation was observed in Col (Figure 8). Collectively, these data show that root NRT2.1 expression in NRT1.1 mutants is blocked in a derepressed state and, thus, that NRT1.1 is required for correct regulation of NRT2.1 by N status of the plant. The same conclusion may also be drawn for the activity of the HATS for NO3–, which appears to be unusually insensitive in chl1-5 plants to the repression exerted by the presence of NH4+ in the nutrient solution (Figures 3 and 7). This role of NRT1.1 in controlling both the regulation of the NO3– HATS activity and NRT2.1 expression seems to be quite specific. First, regulation of AMT1.1 is not affected in chl1-5 plants (Figure 8). Second, the two other known regulations of NRT2.1 expression, namely induction by NO3– (Filleur and Daniel-Vedele, 1999) and regulation by day/night cycle and photosynthates (Lejay et al., 1999, 2003), are also not altered by NRT1.1 mutation (Figure 9).
The mechanism responsible for the role of NRT1.1 in the regulation of NRT2.1 expression is unclear. However, the observation that low NO3– availability in presence of 1 mM NH4+ upregulates NRT2.1 expression in the wild type (Figure 13B) indicates that mutation of NRT1.1 is not strictly required for preventing downregulation of NRT2.1 by N metabolites. NRT1.1 is believed to be the main transporter responsible for NO3– uptake from mixed N sources (Huang et al., 1996; Touraine and Glass, 1997; Crawford and Glass, 1998). Thus, the two situations that lead to overexpression of NRT2.1 in the presence of 1 mM NH4+ (e.g., mutation of NRT1.1 or decrease in external NO3– availability) are both expected to result in a reduced NO3– uptake rate. Accordingly, these situations are associated with lowered NO3– accumulation in tissues (Figures 12 and 13). This strongly suggests that low NO3– uptake rate from mixed NH4NO3 nutrient solution is the actual cause for the upregulation of NRT2.1. Because the presence of 1 mM NH4+ in the medium prevents N deficiency in both wild-type and mutant (see above), this would imply that NRT2.1 expression is specifically repressed by high NO3– uptake rate, independently of the products of NO3– assimilation. Hence, one hypothesis would be that two distinct signaling pathways have to be considered for mediating repression of NRT2.1 by N status of the plant: (1) the well-known feedback repression by N metabolites, related to a specific reduced N status and mediating the reduced N demand for growth of the plant, and (2) a yet unknown feedback repression by NO3– uptake or NO3– content of the tissues, related to the NO3– status of the plant and mediating a specific NO3– demand. The NO3– demand signaling would override feedback repression by N metabolites to stimulate NRT2.1 expression in situations where NO3– uptake rate is low in presence of NH4+ (e.g., in the wild type supplied with nutrient solution at high NH4+/NO3– ratio or in chl1 mutants).
How might NRT2.1 be regulated both by NO3– induction (Filleur and Daniel-Vedele, 1999; Zhuo et al., 1999) and by repression by high NO3– status remains an unanswered question. Opposite direct signaling roles of NO3– (induction/repression) in the regulation of its own uptake systems have already been proposed from physiological studies (Siddiqi et al., 1989; King et al., 1993). However, experiments with NR-deficient mutants or using tungstate, a potent NR inhibitor, provided evidence that on NO3– as sole N source, NRT2.1 is predominantly repressed by products of NO3– assimilation and not by NO3– itself (Krapp et al., 1998; Lejay et al., 1999; Zhuo et al., 1999). On the other hand, it is now well documented that NO3– acts both as a positive and a negative signal for the development of the root system, independently of the reduced N status of the plant (Scheible et al., 1997; Stitt, 1999; Zhang et al., 1999). The model proposed for regulation of lateral root development by NO3– (Zhang et al., 1999; Zhang and Forde, 2000) is of major interest in our context. It postulates repression of lateral root elongation by two separate signaling pathways, one dependent on feedback repression by NO3– itself and the other one dependent on feedback repression by reduced N metabolites (e.g., the same dual mechanism as the one we propose above for regulation of NRT2.1 expression). Furthermore, a local stimulatory effect of NO3– has also been documented for lateral root emergence (Zhang and Forde, 2000), showing that NO3– can indeed play opposite roles on a specific process, depending on the conditions. Clearly, the hypothesis that NRT2.1 expression may be upregulated by NO3– demand through direct repression by NO3– itself deserves particular attention because it creates a strong parallel between the regulatory networks involved in the control of two highly interdependent components of NO3– acquisition by the plant (e.g., the NO3– uptake systems and the size and architecture of the root system).
It is unclear whether the putative NO3– demand signaling triggers upregulation of NRT2.1 only in presence of NH4+ in the external medium or also in other circumstances. For instance, it can also account for the high NRT2.1 transcript level found in the roots of chl1-5 plants supplied with NO3– as the sole N source (Figure 6) because lowered NO3– accumulation in the mutant was also observed in this situation (Figure 12). However, this is in contradiction with the conclusion that NRT2.1 is predominantly repressed by downstream N metabolites when plants are supplied with NO3– as the sole N source (see above). Furthermore, lack of derepression of NRT2.1 by N starvation in the chl1-5 mutant (Figure 8) does not fit well with the hypothesis that mutation of NRT1.1 alters NRT2.1 expression through specific regulation by NO3– status only. In that case, N starvation should still alleviate feedback repression by reduced N status, mediated by N metabolites, and would result in a further stimulation of NRT2.1 expression, which is not observed. Alternatively, the lower NO3– content in roots of chl1 mutants (Figures 12 and 13A) may result in a faster loss of NRT2.1 induction by NO3–, which may then prevent any increase in NRT2.1 transcript level upon transfer of the plants to N-free solution.
Without ruling out the NO3– demand signaling as described above, these considerations suggest that other hypotheses could also be envisaged to account for all effects of NRT1.1 mutation on NRT2.1 expression. In particular, the hypothesis that NRT1.1 plays a direct regulatory role and that its activity generates a signal required for feedback repression of NRT2.1 by N metabolites also has the potential to explain our results. According to this, mutation of NRT1.1 in the chl1 mutants would then prevent feedback repression of NRT2.1 by N metabolites in any situation, thus explaining all NRT2.1 expression data obtained in these mutants, even when NO3– is the sole N source (Figure 6). Furthermore, if not repressed in N replete chl1 plants, NRT2.1 obviously cannot be derepressed by N starvation, which then provides a hypothesis for the lack of increase in NRT2.1 transcript level after transfer of chl1-5 plants to N-free medium (Figure 8). Finally, direct repression of NRT2.1 expression by NRT1.1 activity may also account for the fact that NRT2.1 transcript level is high in wild-type plants supplied with 0.1 mM NO3– plus 1 mM NH4+, whereas it is low when NO3– concentration is increased up to 1 or 10 mM, without modifying that of NH4+ (Figure 13B). Indeed, NRT1.1 is dephosphorylated and functions as a low-affinity transporter in plants under high N provision, whereas it is phosphorylated and has a high affinity for NO3– in N-limited plants (Liu and Tsay, 2003). Although the phosphorylation status of NRT1.1 has not been investigated under our specific experimental conditions, we can hypothesize that the supply of 1 mM NH4+ was sufficient to warrant high N provision to the plants and that, accordingly, NRT1.1 was predominantly in the low-affinity form. If this hypothesis is valid, 0.1 mM NO3– in the external medium would have been too low to allow any significant transport activity of NRT1.1, thus preventing generation of the repressive signal for NRT2.1 expression. By contrast, at 1 or 10 mM external NO3–, the low-affinity form of NRT1.1 is significantly or fully active, which then leads to repression of NRT2.1. Additional evidence further suggests a signaling role of NRT1.1 in NRT2.1 regulation. Indeed, one puzzling aspect of our results is that NRT2.1 expression was dramatically stimulated in the wild type by the increase in external NH4+/NO3– ratio, with only a small decrease in NO3– concentration in roots, and almost no change of this concentration in shoot (Figure 13). Although we cannot exclude a stringent control of NRT2.1 expression by the NO3– demand signaling below a threshold level of NO3– accumulation, this may indicate that it is the sensing of the external NO3– concentration or of the NO3– influx, rather than that of NO3– content of the tissues, which is important for the regulation of NRT2.1 in the presence of NH4+ in the medium. Because this regulation is strongly altered in chl1 mutants, NRT1.1 is thus a good candidate for a NO3– flux and/or a NO3– concentration sensor. It is now firmly established in both yeast and plants that specific membrane proteins have a dual transport and signaling role (Lalonde et al., 1999). In yeast, various aspects of N signaling are related to the sensing activity of such proteins. For instance, the permease homolog SSY1 is involved in the regulation of the expression of amino acid and peptide transporters by the external N source (Didion et al., 1998; Iraqui et al., 1999), and the high-affinity NH4+ transporter MEP2 triggers pseudohyphal growth in conditions of N limitation (Lorenz and Heitman, 1998). Most interestingly, both NRT1.1 and NRT2.1 have been recently proposed to trigger morphological changes in the root system of A. thaliana, which are not explained by the purely nutritional role of these proteins (Guo et al., 2001; Rao et al., 2003). Although our data provide additional circumstantial evidence for a sensing function of NRT1.1, much more direct clues are needed for a definite conclusion on this point. Thus, it is not possible yet to exclude any of the two main hypotheses proposed for explaining the surprising role of NRT1.1 in NRT2.1 regulation. Further analysis of the phenotype of chl1 mutants is being performed to answer this question.
Whatever mechanism is responsible for the upregulation of NRT2.1 by low NO3– uptake in the presence of ample NH4+ supply (NO3– demand signaling or lack of NRT1.1-mediated repression), these hypotheses have a strong physiological significance because the ability of the plant to take up NO3– in presence of NH4+ in the external medium prevents the detrimental effects of pure NH4+ nutrition (Salsac et al., 1987; Volk et al., 1992; von Wirén et al., 2000). Indeed, most herbaceous species achieve highest growth rates on a mixed NH4NO3 N source, whereas supply of NH4+ alone generally results in poor growth and various metabolic disorders (Mehrer and Mohr, 1989; Walch-Liu et al., 2000). Despite its importance, no regulatory mechanism was known to specifically promote NO3– uptake from mixed N source. We suggest here that regulation of NRT2.1 by either NO3– demand or NRT1.1-dependent signaling corresponds to such a mechanism. Thus, in addition to being involved in satisfying the plant's N requirements for growth when NO3– is the only N source (Filleur and Daniel-Vedele, 1999; Lejay et al., 1999; Cerezo et al., 2001; Gansel et al., 2001), NRT2.1 would also play a key role under mixed nutrition in contributing to maintain a healthy balance between NO3– and NH4+ uptake.
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). Briefly, seeds were sown directly on the surface of wet sand in modified 1.5-mL microcentrifuge tubes, with the bottom replaced by a metal screen. The tubes supporting the seeds were placed on polystyrene floating rafts, on the surface of a 10-liter tank filled with tap water. The culture was then performed in a controlled growth chamber with 8-h/16-h day/night cycle at 24°C/20°C. Light intensity during the light period was at 250 µmol m–2 s–1. The basal nutrient solutions supplied to the plants are those described by Gansel et al. (2001) and contained either 1 mM NO3– or 1 mM NH4NO3 as N source. For specific experiments involving response to N deprivation or to various N sources, the NH4NO3 solution was used, in which NH4NO3 was either omitted or replaced by other N sources indicated in the figures. One week after sowing, the tap water was replaced by diluted (1/10) basal medium. After one additional week, the plants were supplied with undiluted nutrient solution until the age of 5 weeks when experiments generally began. The nutrient solution was replaced every week during this period. During the experiments, nutrient solutions were renewed daily and adjusted at pH 5.8. Except when the effect of day/night cycle was investigated, all harvests and measurements were done 5 h into the light period.
Characterization of the Genomic Deletion in chl1-5
Three consecutive steps of PCR were performed on Col-0 or chl1-5 genomic DNA to map the deletion, using 15 primer pairs designed from T28K15 and F12F1 BAC sequences. At the end of this process, the right and left borders of the deletion were mapped with 1-kb accuracy each. Then, two oligonucleotides, delF (5'-TATCCTTCACACACATgCATgAC-3') and delR (5'-AATgCAgTCATgCAgTTTATgCC-3'), with their related genomic sequences separated by 19.4 kb on chromosome 1, were used to amplify the corresponding region in the chl1-5 mutant. As expected, the large 19.4-kb fragment could not be amplified with Col genomic DNA, but a 1.1-kb fragment was amplified with chl1-5 genomic DNA using Pfu polymerase (Promega, Madison, WI). The 1.1-kb fragment obtained was cloned using pCR blunt plasmid (Invitrogen, Carlsbad, CA) and DH5
competent cells. DNA was then sequenced by Genome express (Grenoble, France).
Isolation of chl1-10 and chl1-11 Chlorate-Resistant Mutants
The chl1-10 and chl1-11 chlorate resistant mutants were isolated from the INRA collection of T-DNA insertion lines of Arabidopsis thaliana (ecotype Ws, Versailles, France). The screen was done on 6-d-old seedlings germinated on soil. Chlorate treatment was performed by subirrigating plants every 2 d during 15 d with a nutrient solution containing 2 mM KClO3 and 2 mM NH4NO3 as sole nitrogen source. DNA gel blots performed using probes for right and left borders of the T-DNA suggested that chl1-10 and chl1-11 carry one and four insertions, respectively. An allelism test performed with the chl1-5 mutant indicated that two mutants, named chl1-10 and chl1-11, belong to the chl1 complementation group. DNA gel blots performed using a specific probe for NRT1.1 showed that the structure of this gene was disturbed in both chl1-10 and chl1-11 mutants. The disruption of the NRT1.1 gene in the chl1-10 mutant was characterized by PCR amplification and sequencing of the T-DNA flanking sequences using primers specific to both left and right T-DNA borders (5'-GTCGGCTATTGGTAATAGGA-3' and 5'-CCACAGGCCGTCGAGTTTT-3', respectively) and NRT1.1 flanking genomic sequence (5'-GACGTAGAAGACTGCCATCGATG-3' and 5'-TTTGTCATGCATGTGTGTGAAGG-3', respectively).
SAGE Protocol
The root samples harvested from Col-0 and chl1-5 NH4NO3-grown plants were stored at –80°C before total RNA extraction. The SAGE libraries were obtained from 100 µg of total RNA, using SAGE protocol described by Virlon et al. (1999), with the difference that the anchoring enzyme was MboI (New England Biolabs, Beverly, MA) instead of Sau3AI. Poly(A) RNAs were isolated from 100 µg of total RNA using Dynabeads mRNA direct kit (Dynal, Brown Deer, WI) based on oligo(dT)25 bound covalently to magnetic beads. cDNA were synthesized directly on the beads, and all enzymatic steps needed before digestion by BsmFI were performed on cDNA linked to the beads. All oligonucleotides, with sequences and modifications identical to Virlon et al. (1999), were from Eurobio (Les Ulis, France).
Final concatemers were cloned in pBluescript II KS– from Stratagene (La Jolla, CA), digested by EcoRV, dephosphorylated, and purified on agarose gel. Ligation was performed overnight at 16°C and ElectroMAX DH10B Escherichia coli cells (Life Technologies, Cleveland, OH) were then used for transformation by electroporation. Sequencing was performed as described previously (Fizames et al., 2004) in the Department Genome et Développement des Plantes (University of Perpignan, France) and Genome Express (Grenoble, France). Altogether, 1176 runs of sequencing were needed to obtain the 28,952 tags of the chl1-5 root SAGE library and 1335 runs for the 31,354 tags of the Col-0 root SAGE library.
SAGE Data Analysis
The whole procedure developed to obtain transcript profiles from concatemer sequences is described in Fizames et al. (2004). Briefly, experimental tag sequences were extracted from the concatemer sequences using DIGITAG software (Piquemal et al., 2002). Tag to gene assignment was then performed by matching the sequences of the experimental tags with those of virtual tags extracted from 26,620 annotated genes of the A. thaliana genome (ftp://ftpmips.gsf.de/cress/arabidna/arabi_genomicplus500_v111102.gz), taking into account the coding sequence plus 400 bp 5' and 3' extensions. We have previously determined (Fizames et al., 2004) that this procedure allows the identification of the transcripts corresponding to 60% of the tags found experimentally, with a specificity of 85% (only 15% of the experimental tags match more than one gene), and a reliability of 88% (only 12% of the experimental tags are assigned to wrong genes). The statistical analysis of SAGE data for identification of genes differentially expressed between roots of Col-0 and chl1-5 plants was performed as described in Piquemal et al. (2002).
RNA Extraction and RNA Gel Blot Analysis
Total RNA extraction was performed on roots as described previously (Lobreaux et al., 1992). For RNA gel blot analysis, total RNA (15 µg) was separated by electrophoresis on 3-(N-morpholino)-propanesulfonic acid formaldehyde agarose gel and blotted on nylon membrane (Hybond N+; Amersham Pharmacia Biotech, Buckinghamshire, UK). Membranes were prehybridized for 2 h at 65°C in Church buffer (0.5 M NaHPO4, 1% BSA, and 7% SDS, pH 7.2, with H3PO4). Hybridizations were performed overnight at 65°C after addition of a randomly primed 32P-labeled cDNA probe in the prehybridization buffer. Membranes were washed twice at room temperature for 2 min and twice at 65°C for 15 min with 0.5x SSC (1x SSC is 0.15 M NaCl and 0.015 M sodium citrate) and 0.1% SDS. DNA probes used in this study correspond to full-length cDNAs. A 25S rRNA probe was used as reference for quantification achieved using a PhosphorImager (Storm; Molecular Dynamics, Sunnyvale, CA).
15NO3– and 15NH4+ Uptake
Root 15NO3– or 15NH4+ influxes were assayed as described by Delhon et al. (1995) for NO3– and by Gazzarrini et al. (1999) for NH4+. Briefly, the plants were sequentially transferred to 0.1 mM CaSO4 for 1 min, to complete nutrient solution, pH 5.8, containing either 15NO3– or 15NH4+ (99% atom excess 15N) for 5 min at the concentrations indicated in the figures, and finally to 0.1 mM CaSO4 for 1 min. Roots were then separated from shoots, and the organs dried at 70°C for 48 h. After determination of their dry weight, the samples were analyzed for total N and atom percentage 15N using a continuous-flow isotope ratio mass spectrometer coupled with a C/N elemental analyzer (model ANCA-MS; PDZ Europa, Crewe, UK) as described in Clarkson et al. (1996). Each influx value is the mean of 6 to 12 replicates.
Amino Acid and NO3– Analysis
After harvest of the plants, roots and shoot were separated and stored either at –20°C for amino acid analysis or dried for 48 h at 70°C for NO3– analysis. Free amino acids were extracted from 0.5 g of frozen tissue by grinding in 2 mL of EtOH. The extracts were then left for 1 h at 4°C before centrifugation for 10 min at 2400 rpm and at 4°C. Supernatants were recovered and pellets were subjected to three additional extraction steps identical to the first one, except that these were performed in 1 mL of 80% EtOH, 60% EtOH, and water, respectively. The four supernatants from the same sample were pooled, and an aliquot of this solution filtered (0.45 µm) for amino acid quantification by HPLC (gradient pump SP8800 [Spectra Physics, Mountain View, CA], fluorimeter 821-FP [Jasco, Easton, MD], and SP4270 integrator piloted by SP-LABNET software [Spectra Physics]).
Extraction of NO3– was performed in 0.1 N HCl overnight at 4°C. The NO3– concentration in the extracts was determined colorimetrically at 540 nm after reduction to NO2– on a Cd column and addition of sulfanylamide and N-naphtyl-ethylene-diamine-dichloride.
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Acknowledgments |
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Footnotes |
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The author responsible for distribution of materials integral to the findings presented in this article in accordance with the policy described in the Instructions for Authors (www.plantcell.org) is: Alain Gojon (gojon{at}ensam.inra.fr).
Online version contains Web-only data.
Article, publication date, and citation information can be found at www.plantcell.org/cgi/doi/10.1105/tpc.104.024380.
Received May 17, 2004; accepted June 21, 2004.
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REFERENCES |
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TOPABSTRACTINTRODUCTIONRESULTSDISCUSSIONMETHODSREFERENCES |
|---|
Cerezo, M., Tillard, P., Filleur, S., Muños, S., Daniel-Vedele, F., and Gojon, A. (2001). Major alterations of the regulation of root NO3- uptake are associated with the mutation of Nrt2.1 and Nrt2.2 genes in Arabidopsis. Plant Physiol. 127, 262–271.
Clarkson, D.T., Gojon, A., Saker, L.R., Wiersema, P.K., Purves, J.V., Tillard, P., Arnold, G.M., Paams, A.J.M., Waalburg, W., and Stulen, I. (1996). Nitrate and ammonium influxes in soybean (Glycine max) roots: Direct comparison of 13N and 15N tracing. Plant Cell Environ. 19, 859–868.[CrossRef]
Crawford, N.M., and Glass, A.D.M. (1998). Molecular and physiological aspects of nitrate uptake in plants. Trends Plant Sci. 3, 389–395.[CrossRef][Web of Science]
Delhon, P., Gojon, A., Tillard, P., and Passama, L. (1995). Diurnal regulation of NO3- uptake in soybean plants. I. Changes in NO3- influx, efflux, and N utilization in the plant during the day/night cycle. J. Exp. Bot. 46, 1585–1594.
Didion, T., Regenberg, B., Jorgensen, M.U., Kielland-Brandt, M.C., and Andersen, H.A. (1998). The permease homologue Ssy1p controls the expression of amino acid and peptide transporter genes in Saccharomyces cerevisiae. Mol. Microbiol. 27, 643–650.[CrossRef][Web of Science][Medline]
Doddema, H., and Telkamp, G.P. (1979). Uptake of nitrate by mutants of Arabidopsis thaliana disturbed in uptake or reduction of nitrate. II. Kinetics. Physiol. Plant. 45, 332–338.[CrossRef]
Filleur, S., and Daniel-Vedele, F. (1999). Expression analysis of a high-affinity nitrate transporter isolated from Arabidopsis thaliana by differential display. Planta 207, 461–469.[CrossRef][Web of Science][Medline]
Filleur, S., Dorbe, M.F., Cerezo, M., Orsel, M., Granier, F., Gojon, A., and Daniel-Vedele, F. (2001). An arabidopsis T-DNA mutant affected in Nrt2 genes is impaired in nitrate uptake. FEBS Lett. 489, 220–224.[CrossRef][Web of Science][Medline]
Fizames, C., et al. (2004). The Arabidopsis root transcriptome by serial analysis of gene expression. Gene identification using the genome sequence. Plant Physiol. 134, 67–80.
Forde, B.G. (2000). Nitrate transporters in plants: Structure, function and regulation. Biochim. Biophys. Acta 1465, 219–235.[Medline]
Gansel, X., Muños, S., Tillard, P., and Gojon, A. (2001). Differential regulation of the NO3- and NH4+ transporter genes AtNrt2.1 and AtAmt1.1 in Arabidopsis: Relation with long-distance and local controls by N status of the plant. Plant J. 26, 143–155.[CrossRef][Web of Science][Medline]
Gaymard, F., Pilot, G., Lacombe, B., Bouchez, D., Bruneau, D., Boucherez, J., Michaux-Ferriere, N., Thibaud, J.B., and Sentenac, H. (1998). Identification and disruption of a plant shaker-like outward channel involved in K+ release into the xylem sap. Cell 94, 647–655.[CrossRef][Web of Science][Medline]
Gazzarrini, S., Lejay, L., Gojon, A., Ninnemann, O., Frommer, W.B., and von Wiren, N. (1999). Three functional transporters for constitutive, diurnally regulated, and starvation-induced uptake of ammonium into Arabidopsis roots. Plant Cell 11, 937–948.
Guo, F.Q., Wang, R., Chen, M., and Crawford, N.M. (2001). The Arabidopsis dual-affinity nitrate transporter gene AtNRT1.1 (CHL1) is activated and functions in nascent organ development during vegetative and reproductive growth. Plant Cell 13, 1761–1777.
Guo, F.Q., Young, J., and Crawford, N.M. (2003). The nitrate transporter AtNRT1.1 (CHL1) functions in stomatal opening and contributes to drought stress susceptibility in Arabidopsis. Plant Cell 15, 107–117.
Huang, N.C., Chiang, C.S., Crawford, N.M., and Tsay, Y.F. (1996). CHL1 encodes a component of the low-affinity nitrate uptake system in Arabidopsis and shows cell type-specific expression in roots. Plant Cell 8, 2183–2191.[Abstract]
Huang, N.C., Liu, K.H., Lo, H.J., and Tsay, Y.F. (1999). Cloning and functional characterization of an Arabidopsis nitrate transporter gene that encodes a constitutive component of low-affinity uptake. Plant Cell 11, 1381–1392.
Iraqui, I., Vissers, S., Bernard, F., de Craene, J.O., Boles, E., Urrestarazu, A., and Andre, B. (1999). Amino acid signaling in Saccharomyces cerevisiae: A permease-like sensor of external amino acids and F-Box protein Grr1p are required for transcriptional induction of the AGP1 gene, which encodes a broad-specificity amino acid permease. Mol. Cell. Biol. 19, 989–1001.
Jung, S.H., Lee, J.Y., and Lee, D.H. (2003). Use of SAGE technology to reveal changes in gene expression in Arabidopsis leaves undergoing cold stress. Plant Mol. Biol. 52, 553–567.[CrossRef][Web of Science][Medline]
King, B.J., Siddiqi, M.Y., Ruth, T.J., Warner, R.L., and Glass, A.D.M. (1993). Feedback regulation of nitrate influx in barley roots by nitrate, nitrite, and ammonium. Plant Physiol. 102, 1279–1286.[Abstract]
Krapp, A., Fraisier, V., Scheible, W.R., Quesada, A., Gojon, A., Stitt, M., Caboche, M., and Daniel-Vedele, F. (1998). Expression studies of Nrt2:1Np, a putative high-affinity nitrate transporter: Evidence for its role in nitrate uptake. Plant J. 14, 723–731.[CrossRef][Web of Science]
Lalonde, S., Boles, E., Hellmann, H., Barker, L., Patrick, J.W., Frommer, W.B., and Ward, J.M. (1999). The dual function of sugar carriers. Transport and sugar sensing. Plant Cell 11, 707–726.
Lee, J.Y., and Lee, D.H. (2003). Use of serial analysis of gene expression technology to reveal changes in gene expression in Arabidopsis pollen undergoing cold stress. Plant Physiol. 132, 517–529.
Lejay, L., Gansel, X., Cerezo, M., Tillard, P., Muller, C., Krapp, A., von Wiren, N., Daniel-Vedele, F., and Gojon, A. (2003). Regulation of root ion transporters by photosynthesis: Functional importance and relation with hexokinase. Plant Cell 15, 2218–2232.
Lejay, L., Tillard, P., Lepetit, M., Olive, F., Filleur, S., Daniel-Vedele, F., and Gojon, A. (1999). Molecular and functional regulation of two NO3- uptake systems by N- and C-status of Arabidopsis plants. Plant J. 18, 509–519.[CrossRef][Web of Science][Medline]
Liu, K.H., Huang, C.Y., and Tsay, Y.F. (1999). CHL1 is a dual-affinity nitrate transporter of Arabidopsis involved in multiple phases of nitrate uptake. Plant Cell 11, 865–874.
Liu, K.H., and Tsay, Y.F. (2003). Switching between the two action modes of the dual-affinity nitrate transporter CHL1 by phosphorylation. EMBO J. 22, 1–9.[CrossRef][Web of Science][Medline]
Lobreaux, S., Massenet, O., and Briat, J.F. (1992). Iron induces ferritin synthesis in maize plantlets. Plant Mol. Biol. 19, 563–575.[CrossRef][Web of Science][Medline]
Lorenz, M.C., and Heitman, J. (1998). The MEP2 ammonium permease regulates pseudohyphal differentiation in Saccharomyces cerevisiae. EMBO J. 17, 1236–1247.[CrossRef][Web of Science][Medline]
Lorenz, W.W., and Dean, J.F. (2002). SAGE Profiling and demonstration of differential gene expression along the axial developmental gradient of lignifying xylem in loblolly pine (Pinus taeda). Tree Physiol. 22, 301–310.[Abstract]
Matsumura, H., Nirasawa, S., Kiba, A., Urasaki, N., Saitoh, H., Ito, M., Kawai-Yamada, M., Uchimiya, H., and Terauchi, R. (2003). Overexpression of Bax inhibitor suppresses the fungal elicitor-induced cell death in rice (Oryza sativa L.) cells. Plant J. 33, 425–434.[CrossRef][Web of Science][Medline]
Mehrer, I., and Mohr, H. (1989). Ammonium toxicity: Description of the syndrome in Sinapsis alba and the search for its causation. Physiol. Plant. 77, 545–554.[CrossRef]
Nazoa, P., Vidmar, J.J., Tranbarger, T.J., Mouline, K., Damiani, I., Tillard, P., Zhuo, D., Glass, A.D., and Touraine, B. (2003). Regulation of the nitrate transporter gene AtNRT2.1 in Arabidopsis thaliana: Responses to nitrate, amino acids and developmental stage. Plant Mol. Biol. 52, 689–703.[CrossRef][Web of Science][Medline]
Neuteboom, L.W., Ng, J.M.Y., Kuyper, M., Clijdesdale, O.R., Hooykaas, P.J.J., and van der Zaal, B.J. (1999). Isolation and characterization of cDNA clones corresponding with mRNAs that accumulate during auxin-induced lateral root formation. Plant Mol. Biol. 39, 273–287.[CrossRef][Web of Science][Medline]
Ninnemann, O., Jauniaux, J.C., and Frommer, W.B. (1994). Identification of a high affinity NH4+ transporter from plants. EMBO J. 13, 3464–3471.[Web of Science][Medline]
Okamoto, M., Vidmar, J.J., and Glass, A.D.M. (2003). Regulation of NRT1 and NRT2 gene families of Arabidopsis thaliana: Responses to nitrate provision. Plant Cell Physiol. 44, 304–317.
Oostindiër-Braaksma, F.J., and Feenstra, W.J. (1973). Isolation and characterization of chlorate-resistant mutants of Arabidopsis thaliana. Mutat. Res. 19, 175–185.
Orsel, M., Krapp, A., and Daniel-Vedele, F. (2002). Analysis of the NRT2 nitrate transporter family in Arabidopsis. Structure and gene expression. Plant Physiol. 129, 886–896.
Peterman, T.K., and Goodman, H.M. (1991). The glutamine synthetase gene family of Arabidopsis thaliana: Light-regulation and differential expression in leaves, roots and seeds. Mol. Gen. Genet. 230, 145–154.[CrossRef][Web of Science][Medline]
Piquemal, D., Commes, T., Manchon, L., Lejeune, M., Ferraz, C., Pugnere, D., Demaille, J., Elalouf, J., and Marti, J. (2002). Transcriptome analysis of monocytic leukemia cell differentiation. Genomics 80, 361–371.[CrossRef][Web of Science][Medline]
Rao, H., Little, D.Y., and Malamy, J.E. (2003). A high affinity nitrate transporter regulates lateral root initiation. In Proceedings of the 14th International Conference on Arabidopsis Research. (Madison, WI: University of Wisconsin), p. 250.
Rawat, S.R., Silim, S.N., Kronzucker, H.J., Siddiqi, M.Y., and Glass, A.D. (1999). AtAMT1 gene expression and NH4+ uptake in roots of Arabidopsis thaliana: Evidence for regulation by root glutamine levels. Plant J. 19, 143–152.[CrossRef][Web of Science][Medline]
Salsac, L., Chaillou, S., Morot-Gaudry, J.F., Lesaint, C., and Jolivet, E. (1987). Nitrate and ammonium nutrition in plants. Plant Physiol. Biochem. 25, 805–812.[Web of Science]
Scheible, W.R., Lauerer, M., Schulze, E.D., Caboche, M., and Stitt, M. (1997). Accumulation of nitrate in the shoots acts as a signal to regulate shoot-root allocation in tobacco. Plant J. 11, 671–691.[CrossRef][Web of Science]
Scholten, H.J., and Feenstra, W.J. (1986). Uptake of chlorate and other ions in seedlings of the nitrate-uptake mutant B1 of Arabidopsis thaliana. Physiol. Plant. 66, 265–269.[CrossRef]
Siddiqi, M.Y., Glass, A.D.M., Ruth, T.J., and Fernando, M. (1989). Studies of the regulation of nitrate influx by barley seedlings using 13NO3-. Plant Physiol. 90, 806–813.
Stitt, M. (1999). Nitrate regulation of metabolism and growth. Curr. Opin. Plant Biol. 2, 178–186.[CrossRef][Web of Science][Medline]
Touraine, B., and Glass, A.D. (1997). NO3- and ClO3- fluxes in the chl1-5 mutant of Arabidopsis thaliana. Does the CHL1-5 gene encode a low-affinity NO3- transporter? Plant Physiol. 114, 137–144.[Abstract]
Tsay, Y.F., Schroeder, J.I., Feldmann, K.A., and Crawford, N.M. (1993). The herbicide sensitivity gene CHL1 of Arabidopsis encodes a nitrate-inducible nitrate transporter. Cell 72, 705–713.[CrossRef][Web of Science][Medline]
Velculescu, V.E., Zhang, L., Vogelstein, B., and Kinzler, K.W. (1995). Serial analysis of gene expression. Science 270, 484–487.
Vert, G., Grotz, N., Dedaldechamp, F., Gaymard, F., Guerinot, M.L., Briat, J.F., and Curie, C. (2002). IRT1, an Arabidopsis transporter essential for iron uptake from the soil and for plant growth. Plant Cell 14, 1223–1233.
Virlon, B., Cheval, L., Buhler, J.M., Billon, E., Doucet, A., and Elalouf, J.M. (1999). Serial microanalysis of renal transcriptomes. Proc. Natl. Acad. Sci. USA 96, 15286–15291.
Volk, R.J., Challiou, S., Mariotti, A., and Morot-Gaudry, J.F. (1992). Beneficial effects of concurrent ammonium and nitrate nutrition on the growth of Phaseolus vulgaris: A 15N study. Plant Physiol. Biochem. 30, 487–493.
von Wirén, N., Gazzarini, S., Gojon, A., and Frommer, W.B. (2000). The molecular physiology of ammonium uptake and retrieval. Curr. Opin. Plant Biol. 3, 254–261.[Web of Science][Medline]
Walch-Liu, P., Neumann, G., Bangerth, F., and Engels, C. (2000). Rapid effects of nitrogen form on leaf morphogenesis in tobacco. J. Exp. Bot. 51, 227–237.
Wang, R., Liu, D., and Crawford, N. (1998). The Arabidopsis CHL1 protein plays a major role in high-affinity nitrate uptake. Proc. Natl. Acad. Sci. USA 95, 15134–15139.
Wilkinson, J.Q., and Crawford, N.M. (1993). Identification and characterization of a chlorate-resistant mutant of Arabidopsis thaliana with mutations in both nitrate reductase structural genes NIA1 and NIA2. Mol. Gen. Genet. 239, 289–297.[CrossRef][Web of Science][Medline]
Zhang, H., and Forde, B.G. (2000). Regulation of Arabidopsis root development by nitrate availability. J. Exp. Bot. 51, 51–59.
Zhang, H., Jennings, A., Barlow, P.W., and Forde, B.G. (1999). Dual pathways for regulation of root branching by nitrate. Proc. Natl. Acad. Sci. USA 96, 6529–6534.
Zhuo, D., Okamoto, M., Vidmar, J.J., and Glass, A.D. (1999). Regulation of a putative high-affinity nitrate transporter (Nrt2;1At) in roots of Arabidopsis thaliana. Plant J. 17, 563–568.[CrossRef][Web of Science][Medline]
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