The Arabidopsis Dual-Affinity Nitrate Transporter Gene AtNRT1.1 (CHL1) Is Activated and Functions in Nascent Organ Development during Vegetative and Reproductive Growth

Abstract

The AtNRT1.1 (CHL1) transporter provides a primary mechanism for nitrate uptake in Arabidopsis and is expected to localize to the epidermis and cortex of the mature root, where the bulk of nitrate uptake occurs. Using fusions to GFP/GUS marker genes, we found CHL1 expression concentrated in the tips of primary and lateral roots, with very low signals in the epidermis and cortex. A time-course study showed that CHL1 is activated in the primary root tip early in seedling development and at the earliest stages of lateral root formation. Strong CHL1 expression also was found in shoots, concentrated in young leaves and developing flower buds but not in the shoot meristem. These expression patterns were confirmed by immunolocalization and led us to examine CHL1 function specifically in the growth of developing organs. chl1 mutants showed a reduction in the growth of nascent roots, stems, leaves, and flower buds. The growth of nascent primary roots was inhibited in the mutants even in the absence of added nitrate, whereas elongation of lateral root primordia was inhibited specifically at low nitrate and acidic pH. Interestingly, chl1 mutants also displayed a late-flowering phenotype. These results indicate that CHL1 is activated and functions in the growth of nascent organs in both shoots and roots during vegetative and reproductive growth.

INTRODUCTION

The acquisition of inorganic nutrients involves uptake primarily from the soil solution into roots. Root uptake systems use multiple genes and mechanisms that transport ions from the apoplasm of epidermal and cortical cells into the symplasm. For nitrate uptake, three physiological systems have been described: low affinity, inducible high affinity, and constitutive high affinity (reviewed in Crawford and Glass, 1998; Forde and Clarkson, 1999; Galvan and Fernandez, 2001; Glass et al., 2001; Williams and Miller, 2001). Two families of nitrate transporter genes, NRT1 and NRT2, have been identified that contribute to these uptake systems (reviewed in Crawford and Glass, 1998; Daniel-Vedele et al., 1998; Forde, 2000; Galvan and Fernandez, 2001; Glass et al., 2001; Williams and Miller, 2001). Members of the NRT2 family are part of the high affinity system; NRT2 mutants (Galvan et al., 1996; Filleur et al., 2001) and antisense suppressed lines (Forde, 2000) have reduced high affinity nitrate uptake. NRT2 expression is nitrate inducible; thus, these transporters are part of the inducible high affinity system (reviewed in Forde, 2000; Galvan and Fernandez, 2001; Glass et al., 2001; Williams and Miller, 2001). These genes also display feedback regulation, because ammonium or glutamine, the end products of the nitrate assimilation pathway, suppress NRT2 expression.

The NRT1 family is more complicated and is composed of genes involved in nitrate and amino acid/peptide uptake (reviewed in Crawford and Glass, 1998; Forde, 2000; Galvan and Fernandez, 2001; Glass et al., 2001; Williams and Miller, 2001). The AtNRT1.1 gene (originally named CHL1 in Arabidopsis) was initially described as part of the low affinity nitrate uptake system, because chl1 mutants have reduced low affinity uptake (Doddema and Telkamp, 1979; Tsay et al., 1993; Huang et al., 1996; Touraine and Glass, 1997) and CHL1 in Xenopus oocytes shows a K_m for nitrate in the low affinity (millimolar) range (Tsay et al., 1993; Huang et al., 1996). Subsequent studies revealed that CHL1 also is involved in high affinity nitrate uptake (Wang et al., 1998; Liu et al., 1999) and displays multiphasic uptake kinetics in X. oocytes indicative of a dual-affinity transporter (Liu et al., 1999).

When overall nitrate uptake is examined, CHL1's contribution is found to be dependent on environmental conditions (Huang et al., 1996; Touraine and Glass, 1997; Wang et al., 1998). For example, plants grown without ammonium (i.e., with KNO₃) display little difference in nitrate uptake between the wild type and chl1 mutants; however, with ammonium (i.e., NH₄NO₃), CHL1 can be the predominant transporter in Arabidopsis over a broad range of nitrate concentrations (Touraine and Glass, 1997; Wang et al., 1998). CHL1 is expressed in the absence of nitrate, especially at acidic pH, but it does respond to nitrate with enhanced expression (Tsay et al., 1993; Wang et al., 1998). CHL1 displays little feedback regulation, however (Lejay et al., 1999). These findings indicate that CHL1 functions in multiple uptake systems and is a part of the dynamic response that regulates nitrate uptake in plants.

Because CHL1 plays an integral role in nitrate uptake, one would expect that CHL1 protein would be found predominantly in the outer cell layers of the mature root, where the bulk of nutrient uptake into the symplasm occurs. This proposal is supported by microelectrode experiments showing that nitrate uptake activity, measured as nitrate-induced depolarization across the plasma membranes of root epidermal cells, is reduced by 50% in chl1 mutants (Wang et al., 1998). In addition, analysis of NRT1 localization in tomato showed that transcripts from the nitrate-inducible gene LeNRT1.2 were root-specific and localized to root hairs, whereas those from the constitutively expressed LeNRT1.1 gene were found throughout the root (Lauter et al., 1996). These findings are consistent with the expected localization of CHL1 to epidermal and cortical cells. However, in situ hybridization experiments in Arabidopsis presented an unexpected result. Near the root tip, CHL1 mRNA was found primarily in the epidermis, but farther from the root tip, the mRNA was detected in the endodermis, as if expression becomes more internal in the mature regions of the root body (Huang et al., 1996).

To examine the issue of CHL1 localization in greater detail, translational fusions between CHL1 and GFP/GUS marker genes were made and then examined in transgenic Arabidopsis plants. The results presented here show unexpected patterns of expression, indicating that CHL1 is expressed preferentially in rapidly growing regions in both shoots and roots. Experiments to determine the function of CHL1 using the wild type and chl1 mutants also are described. The phenotypes obtained provide new insights into the function of CHL1 during vegetative and reproductive growth.

RESULTS

CHL1 Is Expressed Preferentially in Growing Regions of Roots and Shoots

To analyze CHL1 expression, transgenic Arabidopsis plants were generated that carried green fluorescent protein (GFP) or β-glucuronidase (GUS) reporter DNAs fused in frame to the CHL1 coding region at two sites: HaeII and XhoI (Figures 1B and 1C). For each construct, >24 transgenic lines were examined. Expression patterns in these lines showed that GFP and GUS signals from the HaeII construct had the same pattern but stronger intensities as those from the XhoI construct (data not shown). For further analysis, transgenic plants from the T2 generation transformed with the HaeII construct were used. Seedlings were grown vertically on plates with 10 mM NH₄NO₃ as nitrogen source, as described in Methods. Plants were examined by confocal microscopy for GFP expression or by light microscopy for GUS staining.

Figure 1.

Schemes of CHL1-GFP/GUS Constructs.

(A) CHL1 gene structure. Closed boxes represent exons containing coding regions, open boxes represent 5′ and 3′ untranslated sequences, and numbers indicate nucleotides from the start of translation.

(B) HaeII reporter construct with ∼4.9 kb of CHL1 promoter sequence and exon 1 (up to amino acid 34 of CHL1) fused in frame at the HaeII site to the coding sequence of GFP or GUS. NOS indicates the 3′ termination sequence of the nopaline synthase gene.

(C) XhoI reporter construct with the 4.9-kb CHL1 promoter and intragenic sequences up to the middle of exon 4 (up to amino acid 269 of CHL1) fused in frame to the coding sequence of GFP or GUS.

When 5-day-old seedlings were examined for CHL1-GFP expression in roots, the strongest fluorescence was found in the tips or growing regions of both the primary root (Figure 2A) and lateral roots (Figure 2E). Tissue staining of the same samples with propidium iodide showed the outlines of the roots and some internal cell layers (Figures 2B and 2F), which could be superimposed on the GFP images (Figures 2C and 2G). Closer examination of the GFP images revealed that all cell layers in the root tip fluoresced strongly, including cells in the lateral and columella root cap (Figure 2L). In the elongation zone, the signal was much diminished. In the mature parts of the root, signals were observed in the stele, but at lower levels than in the tip (Figure 2E). In the epidermis or cortex of the mature regions of the root, no significant difference was observed between transgenic and nontransgenic lines.

Figure 2.

Analysis of CHL1-GFP/GUS Expression in Roots and Shoots.

Transgenic Arabidopsis plants containing the HaeII CHL1-GFP/GUS constructs described in Figure 1 were analyzed by scanning laser confocal microscopy ([A] to [P]) and light microscopy ([Q] to [T]) of GUS-stained tissues.

(A) to (C) Longitudinal section of a primary root of a 5-day-old seedling with just the GFP signal seen as green (A), the image from a propidium iodide (PI)–stained root seen as red (B), and the two images in (A) and (B) merged in a two-channel combination (C).

(D), (H), (L), and (P) GFP images for longitudinal sections of the tip of a primary root in a germinating seedling at 1 day (D), 3 days (H), 5 days (L), and 7 days (P). Bars = 100 μm.

(E) to (G) Longitudinal section of a lateral root of a 7-day-old seedling showing GFP signals (E), PI signals (F), and merged GFP and PI images (G).

(I) to (K) Surface views of emerging leaf and cotyledons at the tip of a 5-day-old seedling showing GFP signals (I), PI signals (J), and merged GFP and PI images (K).

(M) to (O) Longitudinal section of a floral bud at stage 11 showing GFP signals (M), PI signals (N), and merged GFP and PI images (O).

(Q) GUS staining of a young inflorescence within 24 hr of bolting.

(R) GUS staining of isolated floral buds at different stages of development.

(S) GUS staining of an opened floral bud at stage 13.

(T) GUS staining of a 5-day-old seedling showing expression in an immature leaf and stipules.

cot, cotyledon; lp, emerging leaf; pt, petal; se, sepal; sg, stigma; sp, stipules; st, stamen; sy, style.

Next, the time course of CHL1 expression was examined during the first week of seedling growth. Little CHL1 expression was found in seed before germination (data not shown). At day 1, CHL1 expression was activated, with GFP signals highest in the three cell layers of the columella root cap, with weaker signals along the lateral root cap and epidermis (Figure 2D). Almost no expression was detected in the internal layers of the root. At day 3, GFP signals strengthened and expanded into other cell layers, including the root meristematic region (Figure 2H). At day 5, very strong GFP signals were found throughout the root tip (Figure 2L). At day 7, the signal intensities in the root tip began to decrease but continued to extend over the entire tip. These results show that CHL1 expression is activated and expands into the entire root tip during the early stages of seedling growth.

The presence of high GFP signals in the tips of roots suggests that CHL1 expression is targeted to cells in rapidly growing regions. To test this possibility further, growing regions in the shoot were examined using CHL1-GFP and CHL1-GUS marker constructs. Strong GFP signals and GUS staining were observed in emerging and immature leaves (Figures 2I to 2K, 2Q, and 2T) and flower buds (Figures 2M to 2O and 2Q to 2S). Close examination of vegetative shoot tips showed high levels of GFP signal and GUS staining throughout emerging leaves and in young stipules but little signal in mature leaves and, significantly, little to no signal in the shoot apical meristem (Figures 2I and 2T and data not shown). In flower buds, signals increased significantly during bud development (Figure 2R) and then, after fertilization and flower opening, GUS staining decreased (data not shown). GUS and GFP signals were not uniform throughout the buds, because the strongest signals were observed in the reproductive organs (i.e., gynoecium and anthers compared with sepals and petals; Figures 2M and 2S). With short GUS staining times, the strongest signals were observed in the style, stigma, and anthers (Figure 2S). As the flowers aged, GUS staining decreased, retaining some signal in the stigma and pollen grains but with almost no staining in sepals, petals, and carpels (data not shown).

Protein Gel Blot Analysis and Immunolocalization of CHL1 Protein

To verify that the CHL1-GFP/GUS constructs were accurately reporting where CHL1 was expressed, immunolocalization of CHL1 protein was performed. Polyclonal antisera were raised against the 15 N-terminal amino acids of CHL1. To check the specificity of the antisera, affinity-purified antibodies were used to immunostain SDS-PAGE protein blots of microsomal proteins isolated from roots of 6-day-old seedlings. Protein gel blots showed the antibody bound to a protein corresponding to the size of CHL1 (65 kD; Tsay et al., 1993) in microsomal preparations from wild-type plants. This band was missing in microsomal preparations from a chl1 deletion mutant (chl1-5) and in the soluble protein fraction of wild-type plants (Figure 3). These data show that the antibodies specifically recognize CHL1 protein.

Figure 3.

Protein Gel Blot Analysis Using Anti-CHL1 Antibody.

Total protein from roots of 6-day-old seedlings was fractionated into soluble fraction (Sol) and total microsomal membranes (Ms) by differential centrifugation as described in Methods. Samples were separated by 8% SDS-PAGE (10 μg each) and analyzed on protein gel blots. A band at 65 kD was present in Ms lanes from wild-type plants but not in the Ms lane from the chl1-5 mutant or the Sol wild-type lane.

The anti-CHL1 antibody was then used for immunolocalization with whole Arabidopsis seedlings. Images of samples incubated with anti-CHL1 or preimmune sera were visualized by confocal microscopy. The immunofluorescence signals gave patterns very similar to those observed with the GFP/GUS reporter constructs. High levels of immunofluorescence were detected in primary root tips (Figures 4A and 4B), in lateral root tips (Figures 4E and 4F), in emerging leaves (Figures 4I and 4J), in anthers and the gynoecium (Figures 4M and 4N, showing a bud at stage 12), in sepals (Figures 4Q and 4R), and in stipules (Figures 4S and 4T). The second panel in each pair of images shows a composite of immunofluorescence from the antibody staining and the propidium iodide staining. Little signal was detected in tissues treated with preimmune sera (Figures 4C, 4D, 4G, 4H, 4K, 4L, 4O, and 4P).

Figure 4.

Immunolocalization of CHL1 in Roots and Shoots.

Whole Arabidopsis seedlings or organs were incubated with affinity-purified anti-CHL1 antibody for immunocytochemical analysis as described in Methods. Immunofluorescence was detected by confocal microscopy, which shows green signals where CHL1 is present. Organs also were stained with propidium iodide (PI), which emits red light, and a composite image was made containing both CHL1 and PI signals.

(A) to (D) Longitudinal sections of primary roots in 4-day-old seedlings, showing CHL1 immunofluorescence (A), a composite of CHL1 and PI signals (B), a seedling incubated with preimmune serum as a control (C), and a composite of preimmune and PI signals (D).

(E) to (H) Longitudinal sections of lateral roots in 6-day-old seedlings showing CHL1 immunofluorescence (E), a composite of CHL1 immunofluorescence and PI signals (F), a seedling incubated with preimmune serum as a control (G), and a composite of preimmune and PI signals (H).

(I) to (L) Surface views of emerging leaves in shoot tips of 5-day-old seedlings showing CHL1 immunofluorescence (I), a composite of CHL1 and PI signals (J), a seedling incubated with preimmune serum as a control (K), and a composite of preimmune and PI signals (L).

(M) to (P) Longitudinal sections of floral buds showing CHL1 immunofluorescence (M), a composite of CHL1 and PI signals (N), floral buds incubated with preimmune serum as a control (O), and a composite of preimmune and PI signals (P).

(Q) and (R) Surface views of a floral bud cluster showing CHL1 immunofluorescence (Q) and a composite of CHL1 and PI signals (R).

(S) and (T) Surface views of emerging leaves and stipules showing CHL1 immunofluorescence (S) and a composite of CHL1 and PI signals (T).

cot, cotyledon; lp, emerging leaf; g, gynoecium; st, stamen; sp, stipule.

CHL1 Expression during Lateral Root Formation

We next examined CHL1 expression during the initiation and elongation of lateral roots, critical events that establish the architecture of the root. Lateral root initiation begins with the activation of cell division in pericycle cells, which produces a lateral root primordium (Laskowski et al., 1995; Malamy and Benfey, 1997a, 1997b). Primordia emerge from the mature root, forming a new meristem. Eight development stages have been defined during this process (Malamy and Benfey, 1997b). CHL1 expression showed activation at the earliest stage of lateral root initiation (Figures 5A and 5B). Strong CHL1-GFP signals were observed in dividing pericycle cells, signals that were higher than in the stele. These strong signals were maintained during the formation of lateral root primordia and emergence. Strong signals were found in the outer and inner cell layers of the primordium at stages II and III (Figure 5C), in the full primordium at stages IV to VII (Figure 5D), in the emerging primordium (Figure 5E), and in the fully emerged lateral root (Figure 5F). These results show that CHL1 expression is activated at the earliest stages of lateral root formation, well before primordia emerge from the mature root.

Figure 5.

Analysis of CHL1-GFP Expression during Lateral Root Formation.

Images of lateral root primordia were taken using scanning laser confocal microscopy to visualize CHL1-GFP expression. The stages of lateral root development shown are listed below.

(A) Stage I, when cell division just initiates in the pericycle.

(B) Stage I, after several anticlinal divisions in the pericycle.

(C) Stage II, when the first periclinal division has divided the lateral root primordium into two layers, outer and inner.

(D) Stage VI, not yet emerged.

(E) Emerging lateral root primordium.

(F) Fully emerged lateral root with mature tissue pattern.

Bars = 100 μm.

CHL1 Function Is Required for Lateral Root Formation but Not Initiation at Low Nitrate Concentrations

The early activation of CHL1 expression in developing organs of the shoot and root suggests that CHL1 may play a role in nascent organ growth. Our first test of this was to examine the formation and elongation of lateral roots in the wild type (Columbia ecotype) and chl1-5 deletion mutants. We chose conditions in which CHL1 is known to be highly expressed and to make a major contribution to nitrate uptake: growth with NH₄NO₃ at pH 5.5 (Huang et al., 1996; Touraine and Glass, 1997; Wang et al., 1998; Liu et al., 1999). Plants were germinated and grown for 5 days in germination medium (with 10 mM NH₄NO₃, pH 5.5, as described in Methods) and then transferred for 3 days to test medium [germination medium with NH₄NO₃ replaced by 2.5 mM (NH₄)₂SO₄ and 50 μM KNO₃ at pH 5.5]. With these conditions, lateral root length and number were reduced dramatically in chl1-5 plants compared with wild-type plants (Figures 6A, 7B, and 7C); the main root length was little affected (Figure 7A), as was the length of adventitious roots (Figure 6A). An important finding of these experiments is that only the adventitious roots had emerged before the time of transfer to the test medium; no lateral roots were seen.

Figure 6.

Lateral Root Growth in the Wild Type and chl1 Deletion Mutants.

Five-day-old wild-type and chl1-5 seedlings 3 days after transfer from germination medium (10 mM NH₄NO₃, pH 5.5) to test media are shown. All seedlings were grown vertically on agarose plates before and after transfer under continuous light at 24°C. Test media had the same components as the germination medium except that NH₄NO₃ was replaced with the following:

(A) 2.5 mM (NH₄)₂SO₄ and 50 μM KNO₃, pH 5.5.

(B) 2.5 mM (NH₄)₂SO₄ and 50 μM KNO₃, pH 6.5.

(C) 2.5 mM (NH₄)₂SO₄ and 1 mM KNO₃, pH 5.5.

(D) 2.5 mM (NH₄)₂SO₄ and 50 μM KCl, pH 5.5.

Figure 7.

Root Growth and Lateral Root Initiation in Wild-Type and chl1 Mutant Seedlings.

Plants were grown as described in Figure 6. Root length ([A] and [B]) and lateral root number (C) were determined 3 days after transfer to test media. Histograms show the following:

(A) Main root length for each seedling (n = 10).

(B) Sum of lateral root length for each seedling (n = 10).

(C) Number of lateral roots (emerged) for each seedling (n = 20).

(D) Number of lateral roots (emerged), lateral root primordia (not emerged), and the sum of the two for each seedling (n = 20).

Error bars indicate ±sd.

Next, lateral root growth was tested using alternative test media. With low nitrate (50 μM KNO₃) at pH 6.5, lateral root number and length were similar but reduced for wild-type and mutant seedlings (Figures 6B, 7B, and 7C). With higher nitrate concentrations (1 mM KNO₃) at pH 5.5, lateral root number and length showed little difference between wild-type and mutant seedlings (Figures 6C, 7B, and 7C). With no nitrate, lateral root number and length were approximately the same for both wild-type and chl1-5 seedlings (Figures 6D, 7B, and 7C). These results show that CHL1's contribution to lateral root formation and growth is dependent on nitrate concentration and pH. We confirmed these results using another allele of chl1, chl1-1, in the Landsberg background (data not shown).

The reduction in lateral root number observed for the chl1-5 mutant was examined in more detail to determine whether the defect was in lateral root initiation or in a later stage of lateral root formation. The numbers of emerged lateral roots and of primordia that had not emerged were determined by light microscopy. The chl1 mutant has less than half the number of emerged lateral roots than was the wild type (Figure 7D). However, the mutant had more than three times the number of primordia that had not emerged per seedling than the wild type had. When the total number of lateral organs (emerged roots and primordia that had not emerged) was determined, it was found that chl1 and the wild type had approximately the same number of total organs (Figure 7D). Closer examination showed that most of the primordia in the mutant were arrested at stage VII, when the primordia were about to emerge from the primary root. These results indicate that the reduction in lateral root numbers in the mutant is not attributable to a decrease in lateral root initiation but mainly to an inhibition of lateral root primordia maturation or elongation.

CHL1 Function in Primary Root Growth

Another example of nascent organ growth occurs during germination of the seed when the radicle emerges and becomes the primary root. Growth of the primary root was examined for 4 days after germination in wild-type and chl1 mutant seedlings by using two different alleles of chl1 (Figure 8). Under all conditions tested, the primary root grew more slowly during the first 3 days in the mutant but recovered and became almost as long as that in the wild type. This was true at low and high concentrations of nitrate and even in the absence of nitrate (Figure 8). These results are in contrast to what was found for lateral roots, in which the effect of a chl1 mutation was observed only at low nitrate.

Figure 8.

Growth of Primary Roots in Wild-Type and chl1 Plants.

Length of the primary root was determined for seedlings grown with different nitrogen sources for 4 days after germination under continuous light. The wild type and two chl1 mutants are all of the Columbia ecotype. Error bars indicate ±sd.

CHL1 Function in Vegetative and Reproductive Shoot Growth

In the experiments described above, we examined CHL1 function in root development. Because CHL1 also is expressed in the nascent organs of the shoot, it is possible that CHL1 functions in shoot organ development. We examined growth rates of three organs—leaves, primary inflorescence, and flower buds—for plants propagated on peat soil. In all three cases, growth of the nascent organ was reduced in the mutant compared with the wild type for both chl1-5 and chl1-1 alleles. Immature leaves of the mutant expanded more slowly than they did in the wild type, but the fully mature leaves were almost as large (data not shown). When the plants shifted to reproductive development and bolted, the stem of the mutant elongated at a slower rate than did the stem of the wild type (1.4 ± 0.2 cm/day for chl1-5 and 3.0 ± 0.4 cm/day for wild type). After 4 to 5 days, the stem of the mutant elongated at a faster rate, so the final length was about the same as that of the wild type. During floral production, young buds from the mutant expanded at a slower rate than did buds from the wild type (0.4 mm/day for chl1-5 versus 0.65 mm/day for Columbia wild type and 0.25 mm/day for chl1-1 versus 0.6 mm/day for Landsberg wild type), but older buds expanded more rapidly in the mutants (0.55 mm/day for both chl1-5 and chl1-1). These results show that CHL1 function is evident during the early phases of shoot organ growth.

We also examined the effect of chl1 mutations on the overall growth of the roots and shoots in young plants. Fresh weights were measured for wild-type and chl1-5 seedlings germinated and grown on agarose plates under different conditions for 8 days. At 8 days, the chl1 mutants had noticeably smaller leaves, but root number and length were similar to those of the wild-type plants. Fresh weights of the shoot were lower in the mutant in all conditions tested (NO₃⁻ from 100 μM to 10 mM with or without ammonium at pH 5.5 and 6.5; Figure 9). Fresh weights of the roots also were lower in the mutant, but the effect was more dependent on the medium conditions, with no ammonium at pH 5.5 showing the most dramatic effects and ammonium at pH 6.5 showing the least difference (Figure 9). Similar overall results were observed for the chl1-1 allele compared with the parental line in the Landsberg ecotype (data not shown).

Figure 9.

Seedling Growth for Wild-Type and chl1 Mutant Seedlings at Different Nitrogen Conditions.

Plants were grown vertically on agarose plates under continuous light at 24°C for 8 days, and then shoot and root fresh weight for each seedling (n = 10) was determined. The germination media were as described in Methods except that NH₄NO₃ was replaced with the following:

(A) KNO₃ (10 μM to 10 mM), pH 5.5.

(B) 2.5 mM (NH₄)₂SO₄ and KNO₃ (10 μM to 10 mM), pH 5.5.

(C) KNO₃ (10 μM to 10 mM), pH 6.5.

(D) 2.5 mM (NH₄)₂SO₄ and KNO₃ (10 μM to 10 mM), pH 6.5.

Error bars indicate ±sd.

CHL1 Function in Flower Timing

Because chl1 mutants showed reduced stem elongation rates after bolting, we examined flowering time. We expected to see a longer time to the first open flower based on days from germination or number of leaves present. This was observed for both chl1-5 and chl1-1 mutants grown with long days (Figure 10) and 24 hr of light (data not shown). A delay in bolting time also was observed for long day conditions (Figure 10). These results indicate that CHL1 plays an important role in reproductive growth, affecting flower timing and flower bud expansion.

Figure 10.

Flowering Time for Wild-Type and chl1 Mutant Plants.

Histograms show number of days from germination to the initiation of bolting or appearance of the first open flower (A) and number of leaves present per seedling at the time of bolting or the first open flower (B) for plants grown with cycles of 16 hr of light and 8 hr of dark on peat soil. Each measurement was taken from 12 pots with six seedlings each. Col-0, Columbia ecotype; Ler, Landsberg (erecta) ecotype. Error bars indicate ±sd.

DISCUSSION

CHL1 Supports the Growth of Nascent Organs in Roots and Shoots

Our initial prediction, based on the known function of CHL1 and on NRT1 localization studies in tomato, was that CHL1 expression would occur primarily if not exclusively in root epidermal and cortical cells, where the bulk of nitrate uptake occurs. What we discovered was quite different and surprising. Epidermal and cortical cells in mature regions of the root had the weakest expression. The strongest expression was found in the growing tips of the primary and lateral roots. This observation suggested that CHL1 expression is targeted to regions containing rapidly dividing cells. This idea was further supported when lateral roots were examined in detail. At the earliest stages of lateral root formation, such as during the initial cell divisions in the pericycle, an increase in CHL1 expression was observed that was maintained and enhanced as the lateral root primordia developed and emerged from the primary root body. Activation of CHL1 expression also occurred during the growth of the nascent primary root in the first few days after germination. We then examined the shoot, where little CHL1 expression was expected based on previous RNA gel blot analyses (Tsay et al., 1993; Lauter et al., 1996). High levels of expression were found in emerging leaves, stipules, and flower buds. When more mature organs or tissues were examined (mature regions of the root, mature leaves, stems, and flowers), CHL1 expression levels were very low, with the exception of the stele in the root.

These results show that the overall pattern of CHL1 expression (activation or enhancement during nascent organ development) is similar in roots and shoot; however, because of the nature of root and shoot development, there are some differences. In the roots, CHL1 expression remains high in the root tips as the root elongates but is much lower in the mature parts of the root (except in the stele). In the shoot, CHL1 expression is high in young developing leaves and flower buds but then decreases to very low levels in mature organs.

The findings from these localization studies guided us to reexamine CHL1 function. Using our findings from the expression studies, we specifically examined the growth of nascent organs in both shoots and roots and found it to be defective in chl1 mutants. For every organ examined (primary roots, lateral roots, leaves, stems, and flower buds), nascent organ growth was inhibited in the mutant compared with that of the wild type. In the case of lateral root development, we could identify the stage that was being affected by chl1. If we simplify lateral root development into four stages—(1) initiation, (2) primordia formation, (3) elongation of the primordia to an emerged lateral root, and (4) activation of the lateral root meristem with subsequent elongation of the lateral root (Laskowski et al., 1995; Malamy and Benfey, 1997b)—we see that chl1 mutants are defective in the latter stages only. The chl1 mutants have approximately the same number of initiation events and preemergent primordia as do wild-type plants, so stages 1 and 2 appear normal. Progression through stage 3, however, is inhibited, because emerged lateral roots are greatly reduced in number. In addition, emerged lateral roots show no signs of elongation, so stage 4 also may be inhibited. Thus, elongation and maturation of the lateral root primordia are inhibited, whereas initiation and formation of the primordia are not (Figure 7).

An interesting but puzzling feature of our findings is that the most dramatic phenotypes in chl1 mutants did not always correlate with the exact timing of maximal CHL1 expression. In the mutants, growth of the primary root was very apparent 2 to 3 days after germination, yet CHL1 expression peaked at 5 days, when little difference in growth was seen for the mutant and wild type. In even more mature roots (7 days and older), no difference in growth rates was observed between wild-type and chl1 roots as measured by length, even though CHL1 was highly expressed in mature root tips. This was true for the primary root and emerged lateral roots (e.g., the adventitious roots in Figure 6). One explanation for these results could be that other transporters, especially NRT2 and NRT1.2, provide enough uptake activity for the mature root that CHL1 is not needed for normal growth at this stage. No studies on NRT2 spatial expression patterns have been reported for Arabidopsis, but in Nicotiana plumbaginifolia, NRT2 transcripts have been found concentrated in mature root tips and in lateral root primordia by in situ hybridization (Krapp et al., 1998). Transcripts of NRT1.2, a constitutively expressed, low affinity transporter, also are found in epidermal cells of root tips (Huang et al., 1999).

Comparison of CHL1 and CDC, PROLIFERA, and CYCB1 Expression

When one compares the expression of CHL1 with that of genes expressed in actively dividing cells, an overlap is observed. Two such genes, CDC2 and PROLIFERA, are involved in cell cycling and DNA replication, respectively, and show expression in root tips, lateral roots and leaf primordia, apical meristems, and young flower buds (Martinez et al., 1992; Hemerly et al., 1993; Springer et al., 1995, 2000). The overall patterns of expression reported for CDC2 and PROLIFERA are similar to what we found for CHL1. Some striking similarities are (1) PROLIFERA and CHL1 expression in stipules and young leaves, and (2) CDC2 and CHL1 expression in pericycle cells. There also are some differences: (1) strong PROLIFERA and CDC2 expression in the vegetative shoot meristem, where little to no CHL1 expression occurs, and (2) lack of CDC2 expression in the quiescent center of the root meristem, where CHL1 is strongly expressed. CYCB1 expression in roots also is worth comparing because it occurs in actively dividing cells of the root on the basal side of the quiescent center but not in the root cap (Colon-Carmona et al., 1999). In comparison, CHL1 expression also is found in the meristem but includes the quiescent center and the root cap. All of these results indicate that with the exception of the vegetative shoot meristem, CHL1 expression is targeted to cells in actively growing regions in which genes involved in DNA synthesis or cell cycle control are expressed at high levels.

Role of CHL1 in Nascent Organ Growth

So what is CHL1 doing in nascent organs and root tips, and why is it targeted there? Based on what we know about CHL1, one explanation is that CHL1 accelerates the uptake and thus the assimilation of nitrate in situ for growing tissues. The apical region of roots, for example, is known to have a high demand for nitrogen and to support high levels of in situ nitrate reduction (Granato and Raper, 1989, and references therein). However, our plants were provided with ample ammonium, so it is difficult to conclude that a lack of nitrogen is the cause.

Besides being a source of nitrogen, nitrate serves as a metabolite to buffer acidification from ammonium assimilation and as a signal for growth (reviewed in Crawford, 1995; Stitt, 1999; Zhang and Forde, 2000). We attempted to define further the role of CHL1 in the growth of nascent organs by comparing root growth under a variety of nitrate and pH conditions in wild-type and chl1 mutant plants. What we found was that CHL1 function was dependent on nitrate concentration only for lateral root formation. chl1 mutants showed reduced growth of lateral root primordia only at low nitrate concentration and acidic pH. Specific effects of nitrate on lateral root elongation have been described for Arabidopsis (Zhang et al., 1999; Zhang and Forde, 2000). These results are consistent with the idea that the nitrate uptake activity of CHL1 contributes to nitrate signaling or pH homeostasis during lateral root formation. The growth of nascent primary roots, however, was inhibited in the mutants regardless of the nitrate concentration and was even reduced in the absence of nitrate (Figure 8). This result suggests that CHL1's contribution to growth goes beyond the simple uptake of nitrate.

There are several reports that support this idea. Two show that intracellular pH is affected by CHL1 in the absence of nitrate (Meraviglia et al., 1996; Romani et al., 1996). Another report shows that uptake of Cl⁻ and K⁺ can be affected in chl1 mutants (Scholten and Feenstra, 1986), and yet another demonstrates that a similar protein in Brassica napus (BnNRT1) transports both nitrate and histidine (Zhou et al., 1998). Relevant to this issue is the fact that CHL1 is expressed in the absence of nitrate, especially at more acidic pH values (Tsay et al., 1993). We examined the signals from our CHL1-GFP constructs in plants grown without nitrate and found expression in roots with the same cell type–specific pattern seen with nitrate (data not shown). Together, these results suggest that the role of CHL1 may be different in different organs and that, besides the metabolic role of transporting nitrate, CHL1 may contribute (directly or indirectly) to signaling or sensing of nitrate and, potentially, other metabolites. We must caution that we cannot rule out the possibility that small amounts of nitrate contaminated our media and were responsible for the effects we observed in the absence of added nitrate. Further work is needed to fully explain CHL1's function(s) in nascent organ growth.

CHL1 Function in Flowering

An unexpected finding of our work was that CHL1 participates in the transition from vegetative to reproductive growth, because chl1 mutants have a late-flowering phenotype. The delay in flowering was modest, an average of 3 to 4 days to bolting and 3 to 5 days until the first open flower under long day conditions. CHL1 appears to be a newly identified flower- timing gene, because the only reported late-flowering locus to which CHL1 maps is FHA on chromosome 1; however, this locus has been identified as the CRY2 blue light photoreceptor gene (Guo et al., 1998). CHL1 might affect the source-to-sink relationships in the plant, which could affect the diversion of carbon or nitrogen-to-carbon ratios during floral induction. Further work is needed to assess CHL1's role, but it is intriguing that a sole nitrate transporter can have such an effect on bolting and flowering.

METHODS

Plant Material

Plants of Arabidopsis thaliana ecotype Columbia were used in all experiments unless specified otherwise. Seed was surface-sterilized, first in 70% ethanol for 5 min and then in 5% bleach for 15 min, washed with water, and plated on germination medium [pH 5.5; 10 mM NH₄NO₃, 5 mM K₂HPO₄-KH₂PO₄, 2 mM MgSO₄, 1 mM CaCl₂, 0.05 mM FeSO₄-EDTA, 50 μM H₃BO₃, 12 μM MnSO₄·H₂O, 1 μM ZnCl₂, 1 μM CuSO₄·5H₂O, 0.2 μM Na₂MoO₄·2H₂O, 0.5% sucrose, 1 g/L 2-(N-morpholino)ethanesulfonic acid, 1 mg/L thiamine, 100 mg/L inositol, 0.5 mg/L pyridoxine, 0.5 mg/L nicotinic acid, and 8 g/L agarose]. Plates were kept at 4°C for 2 days and then germinated vertically at 24°C under continuous light.

Plant Transformation

To prepare the CHL1-GFP constructs, a 4.9-kb HindIII-HaeII CHL1 fragment (shown in Figure 1B) was ligated with a HaeII-to-XhoI linker to the HindIII and XhoI sites of pBluescript SK+. A 6.1-kb HindIII-XhoI fragment (shown in Figure 1C) was ligated with a XhoI-to-SpeI linker to the HindIII and SpeI sites of pBluescript SK−. CHL1-GFP reporter constructs were generated by making translational fusions of the 4.9-kb HindIII-XhoI fragment and the 6.1-kb HindIII-SpeI fragment into the HindIII-XhoI and HindIII-SpeI sites of p35S-GFP-JFH1 vector, respectively. This green fluorescent protein (GFP) vector was kindly provided by J.F. Harper (Scripps Research Institute, La Jolla, CA) (Hong et al., 1999).

CHL1-GUS constructs were made with translational fusions of the 4.9-kb HindIII-XhoI fragment and the 6.1-kb HindIII-SpeI fragment into the HindIII-SalI and HindIII-XbaI sites of the pBI101.2-GUS vector, respectively (Clontech, Palo Alto, CA).

Transgenic Arabidopsis plants were produced by vacuum infiltrating 4-week-old plants in Agrobacterium tumefaciens culture containing the appropriate construct (Bechtold et al., 1993). Seed from treated plants were collected and screened for kanamycin resistance. Transgenic plants identified in this generation were classified as T1 plants.

CHL1 Antibody and Protein Gel Blotting

Anti-CHL1 rabbit polyclonal antisera were made against the N-terminal peptide of CHL1 (MSLPETKSDDILLDA, with a Cys residue added at the C terminus for coupling) by Genemed Synthesis (South San Francisco, CA). Antisera were purified by antigen affinity chromatography with CHL1 peptide-coupled agarose columns (SulfoLink Kit; Pierce Chemical Co., Rockford, IL). Peptide coupling was performed according to the procedure recommended by the manufacturer. Anti-CHL1 antibody was eluted with 100 mM glycine, pH 2.5, neutralized immediately with 0.1 volume of 1 M Tris-HCl, pH 8.0, and dialyzed against PBS in a Centricon dialyzer (Millipore Corp., Bedford, MA). Protein gel blotting was performed as described (Hong et al., 1999).

Microsomal Membrane Protein Isolation

All manipulations were conducted in a 4°C cold room or on ice with prechilled buffers. Roots from 6-day-old Arabidopsis seedlings were homogenized with 15 mM Tris-HCl, pH 7.8, 250 mM sucrose, 1 mM EDTA, 2 mM DTT, 1 mM phenylmethylsulfonyl fluoride, and 0.6% (w/v) polyvinylpyrrolidone and centrifuged at 10,000g for 15 min. The resultant supernatant was centrifuged at 100,000g for 1 hr to yield the soluble protein fraction (supernatant) and the total membrane fraction (pellet), which was resuspended in homogenization buffer (0.5 mL/10 g starting material).

Histochemical Localization of GUS

Whole seedlings or tissue from kanamycin plants were submerged in staining solution consisting of 25 mM sodium phosphate buffer, pH 7.0, 2 mM 5-bromo-4-chloro-3-indolyl-β-d-glucuronide cyclohexylamine salt, 0.5 mM ferricyanide, 0.5 mM ferrocyanide, and 10 mM EDTA at 37°C for 4 to 10 hr. The β-glucuronidase (GUS) staining solution was removed, and tissue was incubated in 70% ethanol to “bleach” the tissue.

Whole-Mount Immunolocalization

Immunolocalization was performed as described (Müller et al., 1998) with the following modifications. Whole seedlings or floral buds were vacuum infiltrated in a fixative (4% formaldehyde, 50 mM PIPES, 5 mM EGTA, and 5 mM MgSO₄, pH 7.0 [MTSB]) and then incubated in the same fixative for another 2 hr at 20°C. Seedlings and floral buds were subsequently washed three times for 15 min in 0.01% Triton X-100 in MTSB, washed three times in water, incubated in MTSB with 2% driselase, 0.5% cellulase, and 0.025% pectolyase (Sigma), and then shaken vigorously for 15 min. After washing three times with MTSB, seedlings and floral buds were incubated in blocking buffer (5% BSA and 10% goat serum in MTSB) for 2 hr at 20°C, incubated with anti-CHL1 antibodies, and shaken gently overnight at 4°C. The next day, the samples were incubated for another 2 hr at 20°C, washed three times with MTSB for 5 min each, and then incubated with secondary antibodies (1:100 dilution of Alexa Flour 488–conjugated goat antibodies against rabbit IgG; Molecular Probes, Eugene, OR). Before being analyzed by confocal microscopy, samples were washed three times for 5 min each with MTSB and two times with water.

Confocal Microscopy

For GFP and immunolocalization analysis, whole seedlings or organs were counterstained with 10 μg/mL propidium iodide (Sigma) and mounted in water under glass cover slips. GFP fluorescent signal and immunofluorescence were detected using a confocal laser scanning microscope (model MRC-600; Bio-Rad; available at the National Science Foundation–Whitaker Quantitative Imaging and Confocal Microscopy Resource, University of California at San Diego) equipped with a krypton-argon laser and an inverted microscope (Nikon, Tokyo, Japan). GFP images and Alexa Flour 488 fluorescences were collected using a 530 ± 15-nm emission filter, and propidium iodide images were collected using a 620 ± 15-nm emission filter.