Phosphorylation-Mediated Dynamics of Nitrate Transceptor NRT1.1 Regulate Auxin Flux and Nitrate Signaling in Lateral Root Growth^,

The differential phosphorylation of nitrate transceptor NRT1.1 contributes to the modulation of lateral root development through spatiotemporal plasma membrane dynamics and cellular trafficking.

Abstract

The dual-affinity nitrate transceptor NITRATE TRANSPORTER1.1 (NRT1.1) has two modes of transport and signaling, governed by Thr-101 (T101) phosphorylation. NRT1.1 regulates lateral root (LR) development by modulating nitrate-dependent basipetal auxin export and nitrate-mediated signal transduction. Here, using the Arabidopsis (Arabidopsis thaliana) NRT1.1^T101D phosphomimetic and NRT1.1^T101A nonphosphorylatable mutants, we found that the phosphorylation state of NRT1.1 plays a key role in NRT1.1 function during LR development. Single-particle tracking revealed that phosphorylation affected NRT1.1 spatiotemporal dynamics. The phosphomimetic NRT1.1^T101D form showed fast lateral mobility and membrane partitioning that facilitated auxin flux under low-nitrate conditions. By contrast, nonphosphorylatable NRT1.1^T101A showed low lateral mobility and oligomerized at the plasma membrane (PM), where it induced endocytosis via the clathrin-mediated endocytosis and microdomain-mediated endocytosis pathways under high-nitrate conditions. These behaviors promoted LR development by suppressing NRT1.1-controlled auxin transport on the PM and stimulating Ca²⁺-ARABIDOPSIS NITRATE REGULATED1 signaling from the endosome.

Nitrate is the primary nitrogen source in most plants and plants modulate their root system architecture under high-nitrate (HN) and low-nitrate (LN) conditions (Wang et al., 2018b). NITRATE TRANSPORTER1.1 (NRT1.1), also known as NPF6.3 (Léran et al., 2014), is a dual-affinity nitrate transceptor (transporter/receptor) that localizes on the plasma membrane (PM) and functions in nitrate-dependent regulation of root system architecture (Liu et al., 1999; Liu and Tsay, 2003; Ho et al., 2009; Krouk et al., 2010; Léran et al., 2014; Bouguyon et al., 2015), affecting primary root and lateral root (LR) development through multiple pathways (Remans et al., 2006). Notably, the phosphorylation of NRT1.1 at Thr-101 (T101) is crucial for its transport and sensing functions (Liu et al., 1999; Liu and Tsay, 2003; Ho et al., 2009).

In LR development, NRT1.1 functions in nitrate-dependent auxin transport, regulating auxin accumulation and thereby affecting LR growth (Krouk et al., 2010). A signaling network involving Ca²⁺, Ca²⁺-sensor protein kinases (CPKs), and NIN-like proteins (NLPs) is involved in nitrate-regulated LR development via regulation of primary transcription (Remans et al., 2006; Krouk, 2017; Liu et al., 2017). Furthermore, Ca²⁺-ARABIDOPSIS NITRATE REGULATED1 (ANR1), a transcription factor downstream of NRT1.1 and NIN-like protein 7 (NLP7), has been implicated in LR elongation under HN conditions (Gan et al., 2012). Nevertheless, the mechanisms by which the phosphorylation state of NRT1.1 regulates LR development through auxin transport and the Ca²⁺–CPKs–ANR1 signaling pathway have remained unknown.

PM microdomains are enriched in sterols, sphingolipids, and PM-specific proteins, and are involved in regulating the dynamic behavior and assembly states of PM proteins (Bücherl et al., 2017; Cui et al., 2018). The internalization and intracellular trafficking of PM proteins is regulated via control of PM organization and initiation of vesicle transport processes in response to environmental stimuli (Li et al., 2013; Fan et al., 2015). Combining protein labeling techniques and single-particle tracking (SPT) analysis provides a powerful approach to visualize the dynamic behaviors of individual protein molecules with high spatiotemporal resolution (Li et al., 2011; Wang et al., 2013a, 2018a). As an advanced single-molecule technique, fluorescence cross-correlation spectroscopy (FCCS) provides quantitative information on molecular interactions at the single-molecule and nanosecond-timescale levels (Bacia et al., 2006; Wang et al., 2015; Li et al., 2016).

To elucidate the mechanisms underlying the regulation of LR development via phosphorylation of NRT1.1, we investigated the diffusion dynamics and membrane partitioning of NRT1.1 phosphomimetic and nonphosphorylatable mutants in LR cells, allowing us to link data collected at the individual molecule level to auxin transport at the PM. We also analyzed the intracellular trafficking of these NRT1.1 mutants in LR cells to address the possibility that differential phosphorylation of NRT1.1 contributes to the modulation of NRT1.1-mediated signal transduction during LR development.

RESULTS

NRT1.1 Phosphorylation Affects Basipetal Auxin Transport and LR Development

To investigate whether the phosphorylation state of NRT1.1 affects LR development, we measured the density of visible LRs (>0.5 mm) on 8-d–old wild type, chl1-5 (an NRT1.1 null mutant), NRT1.1^T101A/chl1-5 (a nonphosphorylatable T101A mutant, hereafter abbreviated T101A), and NRT1.1^T101D/chl1-5 (a phosphomimetic T101D mutant, hereafter abbreviated as T101D) Arabidopsis (Arabidopsis thaliana) seedlings (Supplemental Fig. S1A). Among seedlings grown in the absence of nitrate or under LN conditions (0.2 mM), the LR density of T101A seedlings was much higher than that of T101D seedlings (Fig. 1, A and B). By contrast, in HN conditions (1 mm), the LR density of the mutants did not significantly differ from that of wild-type plants (Fig. 1, A and B).

Figure 1.

The phosphorylation state of NRT1.1 affects nitrate-regulated auxin transport and LR growth under LN conditions. A, Phenotype of visible LRs (>0.5 mm) in wild-type (WT), chl1-5, T101A, and T101D seedlings grown for 8 d on medium containing different concentrations (0, 0.2, and 1 mm) of NO₃^–. Arrowheads indicate visible LRs. Scale bar = 1 cm. B, Density of visible LRs (>0.5 mm) in wild-type, chl1-5, T101A, and T101D seedlings grown for 8 d on medium containing different concentrations (0, 0.2 and 1 m mm)) of NO₃^–. Data are means ± sd from seven independent experiments (n = 74–91). C and D, Net IAA influx profile (C) and average net auxin flux (D) in yeast strains expressing empty vector (CK), NRT1.1, T101A, and T101D measured by NMT assay. Data are means ± sd (n = 10). E, Basipetal [³H]IAA uptake (counts per minute) in LRs of wild-type, T101A, and T101D seedlings grown for 10 d on medium containing 0.2 and 1 mm of NO₃^–. Data are means ± se from five independent seedlings per treatment. F and H, Confocal images of DR5::GFP and pseudocolor images (blue-green-red palette) showing fluorescence intensity in LRs of T101A and T101D seedlings exposed to 0.2 mm (F) and 1 mm (H) of NO₃^–: LRs initiating primordia (a); LR primordia before emergence (b); newly emerged LRs (c). Scale bars = 20 μm. G and I, Fluorescence intensity (a.u., arbitrary units) of DR5::GFP in LR primordia before emergence of T101A and T101D seedlings exposed to 0.2 mm (G) and 1 mm (I) of NO₃^–. Boxplots represent mean, 25th, and 75th quartiles (n = 6); whiskers represent ± sd. Significant differences are denoted by letters (P < 0.05; Duncan multiple-comparison test) in (B), (D) and (E), and by asterisks (^*P < 0.05, ^***P < 0.001, ns, not significant; Student’s t test) for differences between T101A and T101D in (B), (D), (E), (G), and (I).

To determine whether the phosphorylation state of NRT1.1 affects auxin transport, we generated transformants of the yeast strain YPH499 expressing empty vector (CK, pESC-mRuby-URA), NRT1.1, T101A, or T101D (Supplemental Fig. S1, B and C). The T101D yeast cells showed significantly greater indole-3-acetic acid (IAA) influx than T101A yeast cells by noninvasive microtest technology (NMT) analysis (Fig. 1, C and D). A parallel experiment in planta showed that under LN, LRs of T101D seedlings displayed a 51% increase in [³H]IAA accumulation compared with the T101A seedlings (Fig. 1E; Supplemental Fig. S1D).

To further analyze the effect of NRT1.1 phosphorylation state on auxin accumulation in LRs, we crossed an Arabidopsis line expressing the auxin-inducible DR5::GFP reporter with wild-type, chl1-5, T101A, and T101D plants. The chl1-5 DR5::GFP offspring exhibited strong fluorescence in LRs in response to different concentrations of nitrate (Supplemental Fig. S2, A–F). Consistent with their low auxin transport capacity, the T101A plants displayed strongly enhanced DR5::GFP expression in the primordia and young LRs, whether grown in nitrate-free medium or in LN, as compared to the T101D plants (Fig. 1, F and G; Supplemental Fig. S2, C-–F). However, there was no significant difference in DR5 activity between the T101A and T101D plants in HN conditions (Fig. 1, H and I; Supplemental Fig. S2, A–F). These data indicate that T101A, and by extension, nonphosphorylated wild-type NRT1.1, enhances LR growth in LN by inhibiting basipetal auxin transport, causing the accumulation of auxin in the tips of LRs.

NRT1.1 Phosphorylation Variants Have Different Spatiotemporal PM Dynamics

The spatiotemporal dynamics of PM proteins could control their biological functions (Kusumi et al., 2012). To gain insight into the effect of NRT1.1 phosphorylation on its dynamic behavior, we generated transgenic plants expressing a C-terminal GFP fused to NRT1.1, T101D, or T101A under the control of the NRT1.1 native promoter in the chl1-5 mutant background (Supplemental Fig. S3A). Confocal images (after mannitol-induced plasmolysis) revealed GFP signals mainly on the PM of epidermal cells in LRs (Supplemental Fig. S3, B and C). Gene expression, immunoblot analysis, and LR phenotype assessment confirmed that each transgenic line was functional (Supplemental Figs. S3, D and E and S4; Supplemental Table S1).

Using variable-angle total internal reflection fluorescence microscopy (VA-TIRFM), we found that under LN and HN conditions, spots of T101D-GFP and T101A-GFP localized on the PM and appeared as dispersed diffraction-limited fluorescent spots (Supplemental Fig. S5A). Sequential images showed that the individual particles stayed on the PM for a few seconds and then rapidly disappeared (Supplemental Fig. S5B; Supplemental Videos S1 and S2). SPT analysis revealed that individual T101D-GFP spots had motion trajectories of more than 4 μm within 12 s, whereas T101A-GFP spots were limited to much shorter motion tracks of 1 μm within 6 s (Fig. 2A).

Figure 2.

NRT1.1 phosphorylation variants have different spatiotemporal PM dynamics in the LR cells. A, Motion trajectories of T101D-GFP (0.2 mm of NO₃^–) and T101A-GFP (10 mm of NO₃^–) at the PM. Left and middle, VA-TIRFM images of T101D-GFP and T101A-GFP; green circles indicate the single particles of GFP and colorful fold lines indicate the motion trajectories of GFP spots. Right, typical time-lapse trajectories of T101D-GFP (blue lines) and T101A-GFP (pink lines) at indicated time points. Bar = 2 μm. B to D, Distribution of motion range (B), particle velocity (C), and diffusion coefficients (D) of T101D-GFP (upper) and T101A-GFP (lower) spots at the PM respectively, exposed to 0.2 and 10 mm of NO₃^–. Results are summations of 10 independent experiments each (n = 12,283 for T101D-GFP and 38,560 for T101A-GFP in B; n = 66,398 for T101D-GFP and 49,562 for T101A-GFP in C; n = 1,181 for T101D-GFP and 3,764 for T101A-GFP in D). E, Bubble plots representing differences among T101D-GFP and T101A-GFP for motion ranges, residence time, and particle velocity. Results are summations of 10 independent experiments each (n = 66,398 spots for T101D-GFP and 49,562 spots for T101A-GFP). F, Selected time courses of GFP emissions after background correction showing T101D-GFP one-step bleaching (left) and T101A-GFP two-step bleaching (right) at the PM under 0.2 and 10 mm of NO₃^– treatment respectively. Data are means ± sd from multiple normalized traces depicted in gray (n = 10).

The motion ranges of the spots had a bimodal distribution, with 91.3% of T101D-GFP spots showing long-distance motion and 8.7% showing short-distance motion (Ĝ = 1.05 ± 0.03 μm and 0.30 ± 0.01 μm, respectively; Fig. 2B, upper). For T101A-GFP, the percentage of spots with long-distance motion (Ĝ = 0.82 ± 0.03 μm) was lower, 84.1%, and the percentage with short-distance motion was higher, 15.9% (Ĝ = 0.33 ± 0.01 μm; Fig. 2B, lower; Supplemental Fig. S6A). Moreover, the peak particle velocity of T101A-GFP was significantly lower than that of T101D-GFP (2.89 ± 0.05 μm/s versus 2.99 ± 0.04 μm/s; Fig. 2C; Supplemental Fig. S6B). However, the diffusion coefficient of T101A-GFP was markedly higher than that of T101D-GFP (Ĝ = 9.10 × 10⁻³ ± 0.22 × 10⁻³ μm²/s versus 6.84 × 10⁻³ ± 0.33 × 10⁻³ μm²/s; Fig. 2D; Supplemental Fig. S6C).

To determine the relationships among motion range, residence time, and velocity of T101D-GFP and T101A-GFP particles, we performed a bubble-plot analysis. The distribution range of T101D-GFP spots was more extensive than that of T101A-GFP spots, indicating that T101D-GFP has a longer lifetime and higher particle velocity (Fig. 2E). Taken together, these results imply that phosphorylation of NRT1.1 at T101 leads to long-distance motion and fast particle velocity with reduced diffusion coefficient on the PM of LR cells, whereas nonphosphorylated NRT1.1 has a shorter motion trajectory and lower particle velocity with a higher diffusion coefficient.

The phosphorylation state of NRT1.1 influences its oligomeric state (Tsay, 2014); therefore, we monitored the maximum fluorescence intensity distribution to visualize oligomerization of T101D and T101A. T101D-GFP spots were distributed with a single peak of 2.05507 × 10⁵, whereas T101A-GFP spots exhibited two peaks of 2.64327 × 10⁵ and 3.56615 × 10⁵ by Gaussian fitting, implying that T101D and T101A exist as monomers and mixed monomers/dimers, respectively (Supplemental Fig. S6, D and E). Furthermore, a single-molecule subunit counting assay was used to determine the accurate monomeric/dimeric states of T101D and T101A, respectively. We analyzed the precise ratio of individual T101D-GFP and T101A-GFP cluster photobleaching steps with a progressive idealization and filtering program (Fig. 2F). Approximately 77% of T101D-GFP particles were in monomer form and only 23% were homodimerized under LN. Notably, the proportion of monomers was lower, 54% and the proportion of dimers was increased to 46% for T101A-GFP under HN (Supplemental Fig. S6F). These results suggest that the ratio of dimer to monomer of NRT1.1 on the PM of LR cells is substantially changed by the phosphorylation modification on T101, which might lead to distinct spatiotemporal characters.

Phosphorylation State Affects NRT1.1 Partitioning into PM Microdomains under Low Nitrate

To assess the relationship between NRT1.1 phosphorylation state and its localization in membrane microdomains, we transiently coexpressed NRT1.1-GFP, T101D-GFP, and T101A-GFP with the fluorescent microdomain reporter AtRem1.3-mCherry in Nicotiana benthamiana leaf epidermal cells (Supplemental Fig. S7). Based on Pearson and Manders coefficient analysis, we found moderately lower colocalization between NRT1.1-GFP and Rem1.3-mCherry under HN compared with LN conditions (Fig. 3, A and B). However, T101D-GFP and AtRem1.3-mCherry exhibited higher Pearson and Manders correlation coefficients, compared with T101A-GFP and AtRem1.3-mCherry, under both HN and LN (Fig. 3, A and B).

Figure 3.

Phosphorylation state affects NRT1.1 partitioning into PM microdomains. A and B, Quantitative colocalization analysis for NRT1.1-GFP, T101D-GFP, and T101A-GFP respectively, with AtRem1.3-mCherry in epidermal leaf cells of N. benthamiana by the Pearson (A) and Manders (B) coefficients. Data are means ± sd (n = 16–21). C, Lifetime maps of the LR cells expressing T101D-GFP and T101A-GFP, with or without AtRem1.3-mCherry exposed to 0.2 and 10 mm of NO₃^–. The pseudocolor scale varied from 1.8 ns to 2.8 ns. Scale bar = 5 μm. D, Average fluorescence lifetime and FRET efficiency of T101D-GFP and T101A-GFP in the presence or absence of AtREM1.3-mCherry. Data are means ± sd (n = 12–16). Significant differences are denoted by different letters (P < 0.05; Duncan multiple-comparisons test) in (A), and by asterisks (^*P < 0.05, ^**P < 0.01, ^***P < 0.001; Student’s t test) for differences between T101D-GFP and T101A-GFP in (A), (B), and (D).

To obtain a more dynamic picture of the colocalization of NRT1.1, T101D, T101A, and AtRem1.3 in epidermal cells of LRs, we used Förster resonance energy transfer-fluorescence lifetime imaging microscopy (FRET-FLIM) on the transgenic lines expressing NRT1.1/T101D/T101A-GFP with AtRem1.3-mCherry (Fig. 3C). In plants coexpressing T101D-GFP with AtRem1.3-mCherry, fluorescence lifetime sharply decreased and the FRET efficiency (τ) increased to 38.7% upon LN treatment and to 18.2% upon HN treatment, in comparison with T101D-GFP alone (Fig. 3D). In contrast, no obvious changes were observed in the fluorescence lifetime of T101A-GFP coexpressed with AtRem1.3-mCherry in comparison with T101A-GFP alone. FRET efficiency between T101A-GFP and AtRem1.3-mCherry was only 0.57% and 2.6% under LN and HN conditions, respectively (Fig. 3D). Collectively, these analyses reveal that PM microdomains contribute to the partitioning of the phosphorylated form of NRT1.1.

Phosphorylation State Affects Secretion and Internalization of NRT1.1

Most PM proteins undergo intracellular trafficking between the PM and endosomal compartments (Wu et al., 2014), and their phosphorylation states are vital elements in the regulation of this trafficking (Offringa and Huang, 2013). To dissect the effects of phosphorylation on NRT1.1 secretion in the epidermal cell of LR, we first carried out a fluorescence recovery after photobleaching assay with NRT1.1, T101D, and T101A. We divided the LR into three bleaching sectors (top, middle, and bottom). If the sectors show differences in recovery, this indicates that the new, unbleached proteins arrive by lateral diffusion; if they show no difference, this indicates the proteins arrive by secretion. Indeed, these sections showed no difference in recovery rate between the three proteins, demonstrating that all three were recruited mainly through secretion rather than lateral diffusion (Supplemental Fig. S8, A and B). The fluorescence recovery rate in T101D-GFP seedlings was much lower than that of T101A-GFP under LN (0.3% for T101D-GFP versus 18.9% for T101A-GFP) or HN conditions (12.7% for T101D-GFP and 49.5% for T101A-GFP; Fig. 4A). Furthermore, brefeldin A (BFA), an inhibitor of vesicle recycling through ADP-ribosylation factor-guanine exchange factors (ARF-GEFs), inhibited fluorescence recovery in T101D-GFP markedly, by 69%, but had a much weaker effect on T101A-GFP, causing only a 15% decrease (Fig. 4A, inset; Supplemental Fig. S8C). These results imply that T101D relies on the ARF-GEFs pathway for effective secretion, whereas T101A does not.

Figure 4.

Phosphorylation state of NRT1.1 influences its exocytosis and endocytosis in the LR cells. A, Fluorescence recovery curves of the photobleached regions in the LR cells expressing T101D-GFP and T101A-GFP, exposed to 0.2 and 10 mm of NO₃^– with or without BFA pretreatment (n = 4). Inset, percentage of decreased fluorescence recovery rate under BFA pretreatment. Asterisks indicate significance (^***P < 0.001; Student’s t test). B to D, Confocal images of NRT1.1-GFP, T101D-GFP, and T101A-GFP in LR cells respectively, exposed to different concentrations (0.2, 1, and 10 mm) of NO₃^– with CHX (B), BFA (C), or CHX + BFA (D) pretreatment. Scale bars = 10 μm. E, Representative time-lapse image series of individual T101D-GFP and T101A-GFP spot in LR cells by VA-TIRFM. Images were collected at 3 Hz; for brevity of presentation, one in six frames (at 2-s intervals) are shown. Scale bar = 300 nm. F and G, Normalized fluorescence intensity (ΔF/F; n = 3) of T101D-GFP (F) and T101A-GFP (G) in LR cells, with ± sd from multiple events in gray. H, Kymograph analysis of T101A-GFP and T101D-GFP fluorescence in LR cells showing individual points from yellow rectangles. Scale bar = 5 s. I, Average lifetime of T101D-GFP and T101A-GFP clusters in LR cells. Boxplots represent mean, 25th and 75th quartiles. Whiskers represent minimum and maximum. n = 6,697 dots are pooled across seven independent experiments for T101D-GFP, and n = 44,066 dots are pooled across nine independent experiments for T101A-GFP. Asterisks indicate significant differences between T101D-GFP and T101A-GFP (^***P ≤ 0.0001, Student’s t test). J, Frequency distribution analysis of T101D-GFP and T101A-GFP lifetimes shown in (I).

Next, we dissected the internalization of NRT1.1, T101D, and T101A in response to different nitrate concentrations in the epidermal cell of LR. After pretreatment with the protein synthesis inhibitor cycloheximide (CHX), the internalization of NRT1.1-GFP, as reflected by the number of fluorescent spots in the cytoplasm, showed a dose-dependent increase after treatment with nitrate at 0.2, 1, and 10 mm (Fig. 4B; Supplemental Fig. S8D). Notably, T101D-GFP was found at the cell surface and was mostly absent from intracellular compartments, whereas T101A-GFP showed substantial accumulation in the cytoplasm (Fig. 4B). Statistical analysis of the internalization spots on the cells of T101A-GFP and T101D-GFP seedlings showed a significantly higher number of spots in T101A-GFP compared with T101D-GFP under both LN (0.2 mm) and HN (10 mm) conditions (Supplemental Fig. S8E).

In parallel experiments to detect the formation of BFA-induced compartments, we found that NRT1.1, T101D, and T101A all clearly accumulated in BFA bodies after treatment with BFA only (Fig. 4C). After treatment with BFA and CHX, T101D-GFP signals were still trapped in BFA compartments, whereas T101A-GFP formed dispersed spots centered around BFA compartments that were also dyed by FM4-64 (a PM-specific dye), a pattern similar to that seen for NRT1.1-GFP after HN treatment (Fig. 4D; Supplemental Fig. S8E). These observations suggest that the absence of phosphorylation on NRT1.1 facilitates its internalization independently of ARF-GEFs.

We then implemented TIRFM analysis to determine the PM residence time of T101D-GFP and T101A-GFP. We used MATLAB (https://www.mathworks.com/) to track 6,696 spots of T101D-GFP and 44,065 spots of T101A-GFP under LN and HN conditions, respectively. T101D-GFP particles stayed on the PM for ∼15 s, whereas T101A-GFP particles had a PM residence time of only 10 s (Fig. 4E). In accordance with this, normalized fluorescence intensity and kymograph tests showed a significantly longer PM lifetime for T101D-GFP than for T101A-GFP (Fig. 4, F–H). Similarly, the residence times of T101D-GFP and T101A-GFP on the PM averaged 6.73 s and 5.63 s, respectively (Fig. 4I).

To obtain more data on PM residence time of T101D-GFP and T101A-GFP, we conducted frequency distribution analysis of lifetime. Almost 53% of T101D-GFP spots persisted for more than 5 s on the PM, whereas over 70% of the T101A-GFP spots resided there for less than 5 s (Fig. 4J). Together, these results reveal that the phosphomimetic mutant of NRT1.1 undergoes slower constitutive endocytosis, whereas the nonphosphorylated form is internalized via a faster inducible endocytosis process.

NRT1.1 Phosphorylation State Dictates Endocytic and Intracellular Trafficking Routes

Endocytosis of PM proteins can be generally categorized into clathrin-mediated endocytosis (CME) and clathrin-independent endocytosis (CIE) pathways (Mayor and Pagano, 2007; Fan et al., 2013, 2015). To gain insight into the mechanism of NRT1.1 internalization, we characterized NRT1.1 endocytosis in the LR cells under the CLATHRIN H CHAIN2 (CHC2) knockout mutant chc2-1 background. The intracellular accumulation of T101D-GFP and T101A-GFP clusters was significantly inhibited in response to LN and HN treatment, indicating that clathrin is required for NRT1.1 endocytosis (Fig. 5, A and B). Quantification of NRT1.1 spots in endocytic vesicles showed that endocytosis was blocked almost totally, by 88.5%, in T101D-GFP/chc2-1 seedlings, but by only 65.2% in T101A-GFP/chc2-1 seedlings (Supplemental Fig. S9A), implying that the CIE pathway may contribute more (22.2%) to the endocytosis of T101A than to that of T101D (Supplemental Fig. S9B). To further test this, we applied methyl-β-cyclodextrin (MβCD), a well-established sterol-interfering agent that is routinely used to block the CIE pathway, to T101A-GFP seedlings. This resulted in a 17.5% decrease in the average number of endocytosis spots (Supplemental Fig. S9, C and D), consistent with the earlier results.

Figure 5.

NRT1.1 phosphorylation state dictates endocytic pathways in LR cells. A, Confocal images showing internalization of T101D-GFP and T101A-GFP either in the chc2-1 mutant background (right) or not in the mutant background (left), exposed to 0.2 mm and 10 mm of NO₃^– with CHX pretreatment. Scale bars = 10 μm. B, Quantification of internalized spots shown in (A). Boxplots represent mean, 25th, and 75th quartiles. The colored points represent 10 independent experiments (n = 254 and 249 for T101D-GFP and T101D-GFP/chc2-1, n = 75 and 86 for T101A-GFP and T101A-GFP/chc2-1, respectively). Whiskers represent ± sd. Asterisks denote statistically significant differences (^***P < 0.001; Student’s t test). C, Confocal images showing colocalization of T101A-GFP with AtCLC-mCherry, AtFlot1-mCherry, and AtRem1.3-mCherry in LR cells, exposed to 10 mm NO₃^–. Scale bar = 1 μm. D, Quantitative colocalization analysis for the data from (C) by Pearson correlation coefficients. The colored points represent three independent experiments (n = 12). Whiskers represent ± sd. Asterisks denote statistically significant differences (^*P < 0.01, ^***P < 0.001; ns, not significant; two-tailed Student’s t test). E to G, FCCS analysis of the fluorescence cross-correlation curves (G[τ]) between T101A-GFP and AtCLC-mCherry (E), AtFlot1-mCherry (F), or AtRem1.3-mCherry (G) in LR cells exposed to 10 mm of NO₃^–. H, Average relative cross correlation of T101A with AtCLC/AtFlot1/AtRem 1.3. Data are means ± sd (n = 75) from at least five independent seedlings per treatment.

We further examined the relationship between T101A and the CME and microdomain marker proteins by using dual-color confocal microscopy in transgenic lines coexpressing T101A-GFP and AtCLC-mCherry, AtFlot1-mCherry, or AtRem1.3-mCherry (Fig. 5C). T101A-GFP populations showed good colocalization with AtCLC-mCherry and AtFlot1-mCherry (Pearson correlation coefficients (r) = 0.123 and 0.152, respectively) but not with AtRem1.3 (r = 0.004; Fig. 5D). By FCCS, T101A-GFP showed high relative cross-correlation amplitude with AtCLC-mCherry and AtFlot1-mCherry (r = 0.88 ± 0.001 and 0.81 ± 0.082, respectively), whereas the fluorescence of T101A-GFP and AtRem1.3-mCherry gave a low relative cross-correlation amplitude of 0.59 ± 0.014 (Fig. 5, E–H). These results indicate that the endocytosis of T101A occurs via both the CME and CIE pathways, whereas T101D is internalized mainly via the CME pathway.

To gain a better understanding of the intracellular trafficking routes of NRT1.1, we investigated the influence of the vacuolar H⁺-ATPase inhibitor concanamycin A (Conc A) on the subcellular localization of NRT1.1 in LR cells, T101A-GFP and T101D-GFP. NRT1.1-GFP and T101A-GFP showed vacuolar targeting after 6 h of treatment, but not within 2 h (Fig. 6A). Consistent with this, 6 h of darkness also led to vacuole-like accumulation of NRT1.1 and T101A-GFP signals. T101D-GFP did not localize to vacuoles under any of the treatments tested. In parallel, we applied the phosphatidylinositol-3-kinase inhibitor wortmannin (WM) together with Conc A to block the recycling of vacuolar sorting receptors between the prevacuolar compartment (PVC) and trans-Golgi network (TGN; Kleine-Vehn et al., 2008). WM strongly reduced vacuolar trafficking of NRT1.1-GFP and T101A-GFP, and the fluorescent proteins accumulated in the typical Conc A bodies (Fig. 6A).

Figure 6.

NRT1.1 phosphorylation state dictates intracellular trafficking routes in LR cells. A, Confocal images of NRT1.1-GFP, T101A-GFP, and T101D-GFP in LR cells, exposed to 10 mm NO₃^– with Conc A for 2 h or Conc A, dark and Conc A plus WM for 6 h. Scale bar = 20 μm. B, Confocal images and quantitative analysis of colocalization between T101A-GFP with TGN, MVB, and LE markers in LR cells respectively, exposed to 10 mm NO₃^–. Arrowheads and arrows point to T101A-GFP vesicles that colocalized and did not colocalize with markers, respectively. Scale bars = 5 μm. Graphs at right show the percentages of T101A-GFP-positive vesicles colocalized with the TGN, MVB, and LE markers respectively (n = 64, 41, 179, and 133, where n is the number of T101A-GFP-positive vesicles). C, Confocal images of colocalization between T101A-GFP with the PVC and vacuole markers respectively, exposed to 10 mm of NO₃^– for 2 h and 6 h. Arrows point to T101A-GFP-positive vesicles that colocalized with markers. Scale bars = 5 μm. D, model for intracellular trafficking between PM and cytoplasm of NRT1.1 phosphorylation variants. Under LN conditions, phosphorylated NRT1.1 presents a constitutive secretion and endocytosis between PM and TGN/EE, which is dependent on the activity of ARF-GEF, whereas under HN conditions, the induced secretion and endocytosis of dephosphorylated NRT1.1 between PM and MVB/PVC compartments increase, which is largely independent of the activity of ARF-GEFs. The gray and pink/blue numbers represent the unknown and quantitative percentage of secretion/endocytosis in total. The numbers in red indicate increased percentage of secretion and endocytosis in total.

Considering that T101D-GFP did not accumulate in vacuoles even under Conc A treatment, we assessed the spatial relationship between T101A-GFP and different fluorescence-tagged endomembrane markers by quantitative colocalization analysis. After 2 h of treatment with HN, T101A-GFP partially colocalized with all tested fluorescent TGN and early endosome (EE) markers with low Pearson correlation coefficients, including VTI12-mRFP (18.8%), VHA-a1-mRFP (19.5%), and CLC-mCherry (5.7%; Fig. 6B). In contrast, T101A-GFP colocalized with 64.8% of the multivesicular body (MVB) marker AtSNX1-mRFP spots (Fig. 6B). Moreover, T101A-GFP signals were well colocalized with the PVC and late endosome (LE) markers (RabG3f-mRFP and mRFP-SYP22) and were delimited within the tonoplast after 2 h and 6 h of Conc A treatment (Fig. 6C). Based on the results of intracellular trafficking of NRT1.1, we propose a model for the transfer of unphosphorylated NRT1.1 between the PM and cytoplasm where unphosphorylated NRT1.1 follows an intracellular trafficking route from the PM to the vacuole, passing through the MVB and PVC compartments, which is largely independent of the TGN and EEs (Fig. 6D).

To clarify whether T101A is also degraded by the proteasome pathway, we treated T101A-GFP seedlings with the specific 26S proteasome inhibitor MG132. Without MG132, no detectable degradation of T101A was observed after 15 min of HN treatment; however, there was a 50% reduction in T101A after 4 h of HN (Supplemental Fig. S10A). Notably, treatment with MG132 reduced T101A to 30% after 4 h of HN, revealing that the 26S proteasome is involved in T101A degradation. To determine whether NRT1.1 was involved in initiating autophagy after nitrogen deficiency, we assessed the colocalization of T101D-GFP and the fluorescent autophagosome marker ATG8-CFP. We observed no colocalization between T101D and ATG8 under nitrogen-rich conditions (Supplemental Fig. S10B), but a clear colocalization under nitrogen-free conditions, as indicated by line-scan analysis (Supplemental Fig. S10C). These results suggest that the proteasome and autophagy pathways provide different mechanisms for NRT1.1 degradation in different nitrate conditions.

Internalization of NRT1.1 Promotes Nitrate-Induced Ca²⁺-ANR1 Signaling and LR Development

Nitrate triggers unique Ca²⁺–CPKs–NLPs signaling in the central nutrient-growth network (Krapp et al., 2014). As a downstream component of NLP7, ANR1 acts as a key regulator of LR proliferation (Mounier et al., 2014). To uncover whether NRT1.1 phosphorylation state affects cytoplasmic calcium concentration ([Ca²⁺]_cyt), we assessed nitrate-triggered Ca²⁺ signaling in the epidermal cell of LR from the various genotypes using Fluo-4 AM dye (Supplemental Fig. S11A). Based on the pseudocolor and kymograph images of wild-type samples, we proposed 60 s as a suitable length of time to monitor [Ca²⁺]_cyt signaling after nitrate stimulation (Supplemental Fig. S11, B–E). As described previously in Riveras et al. (2015), we found that nitrate specifically stimulated an increased Ca²⁺ signature in wild type but not chl1-5 mutant seedlings (Fig. 7A). Under both LN and HN conditions, T101A seedlings showed a transient increase in [Ca²⁺]_cyt, whereas T101D seedlings showed a reduction (Fig. 7B). In parallel to [Ca²⁺]_cyt accumulation, HN-induced expression of ANR1 in LRs was detected in T101A but not in T101D (Fig. 7C). Based on these findings, we conclude that the nonphosphorylated form of NRT1.1 could activate the Ca²⁺–CPKs–NLPs signaling pathway to elevate the expression of ANR1.

Figure 7.

Phosphorylation state of NRT1.1 affects nitrate-regulated [Ca²⁺]_cyt signaling and ANR1 expression. A, Pseudocolor images (blue-yellow-red palette) showing [Ca²⁺]_cyt signals in LR meristematic zone of wild-type (WT), chl1-5, T101D, and T101A seedlings treated with different concentrations (0, 0.2, and 10 mm) of NO₃^– for 60 s. Scale bar = 20 μm. B, Quantification analysis of [Ca²⁺]_cyt signals (a.u., arbitrary units) from (A). The colored points represent three independent seedlings per treatment (n = 30–70). Whiskers represent ± sd. C, Relative ANR1 expression level in LRs of wild-type, chl1-5, T101D, and T101A seedlings treated with different concentrations (0, 0.2, and 10 mm) of NO₃^– for 15 min. Data are means ± se from three independent experiments (n = 22). In (B) and (C), statistically significant differences are denoted by letters (P < 0.05; Duncan multiple-comparisons test), and by asterisks for differences between T101D and T101A (^*P < 0.05, ^***P < 0.001; two-tailed heteroscedastic Student’s t test).

To address the function of internalization in signaling mediated by the nonphosphorylated form of NRT1.1, we first perturbed the internalization pathway and looked for changes in nitrate-triggered [Ca²⁺]_cyt signaling. When T101A seedlings were treated with MβCD, nitrate-induced [Ca²⁺]_cyt accumulation was significantly impaired. Similar results were obtained in T101A/chc2-1 seedlings. Nevertheless, both Conc A and Bafilomycin A1 (Baf A1) treatment promoted [Ca²⁺]_cyt influx (Fig. 8, A and B). We then monitored the expression of NRT1.1 and of ANR1 in LRs, the target of the nitrate-Ca²⁺ signaling cascade. Clathrin mutation and MβCD treatment reduced the induction of both genes, whereas Conc A and Baf A1 treatment upregulated the expression level of ANR1 but not NRT1.1, which can be explained by feedback repression acting through the accumulation of NRT1.1 (Fig. 8, C–F). Moreover, we found that T101A-GFP/chc2-1 seedlings showed limited development of LRs compared with T101A-GFP when grown on HN medium (Fig. 8, G and H). Therefore, the endocytosis of nonphosphorylated NRT1.1 appears to be positively associated with nitrate-triggered signaling.

Figure 8.

LR development and PNR signaling are enhanced by HN-induced endocytosis of NRT1.1. A, Pseudocolor images (blue-yellow-red palette) showing [Ca²⁺]_cyt signals in LR meristematic zone of T101A seedlings grown in NO₃^–-free medium, exposed to 10 mm of NO₃^– with no inhibitors (CK), MβCD, Conc A, or Baf A1 pretreatment, as well as in chc2-1 mutant background. Bar = 20 μm. B, Quantification analysis of [Ca²⁺]_cyt signals (a.u., arbitrary units) from (A). The colored points represent three independent seedlings per treatment (n = 40–60). Whiskers represent ± se. Significant differences are denoted by letters (P < 0.05; one-way ANOVA with Tukey multiple-comparisons test). C and D, Gene expression level of NRT1.1 and ANR1 (C) and protein expression level of NRT1.1 (D) in LRs of T101A-GFP and T101A-GFP/chc2-1 seedlings treated with 10 mm of NO₃^– for 15 min. E and F, Gene expression levels of NRT1.1 and ANR1 (E) and protein expression level (F) of NRT1.1 in LRs of T101A-GFP seedlings exposed to 0 or 10 mm of NO₃^– with no inhibitor, MβCD, Conc A, and Baf A1 (only for E) pretreatment. In (C) and (E), boxplots represent average, 25th and 75th percentiles; colored points represent three independent experiments (n = 9–24); whiskers represent ± sd. Significant differences are denoted by asterisks (^***P < 0.001; Student’s t test). In (D) and (F), staining with Coomassie Brilliant Blue serves as the protein loading control. Numbers refer to quantitative results by the software ImageJ. G and H, LR phenotype (G) and density of LRs (H) in T101A-GFP and T101A-GFP/chc2-1 seedlings grown on 10 mm of NO₃^– medium for 8 d. In (G), arrowheads indicate visible LRs. Scale bar = 1 cm. In (H), boxplots represent average, 25th and 75th percentiles; colored points represent four independent experiments (n = 38 [green] and 51 [gray]); whiskers represent ± sd. Significant differences are at ^***P < 0.001 from Student’s t test.

DISCUSSION

Nitrate has a strong effect on plant morphogenesis and root system architecture (Wang et al., 2018b). The nitrate transceptor NRT1.1 functions as a dual-affinity transporter and nitrate sensor in response to variations in nitrate concentration by switching between phosphorylated and dephosphorylated forms (Liu et al., 1999; Liu and Tsay, 2003; Ho et al., 2009; Léran et al., 2014). Aside from the transceptor function mentioned above, NRT1.1 could transport auxin basipetally via a nitrate-dependent pathway, and trigger the Ca²⁺–CPKs–NLPs signaling pathway to regulate LR development (Krouk et al., 2010; Liu et al., 2017). However, little is known about how different NRT1.1 phosphorylation variants may be associated with the regulation of LR growth. Here, we provide several lines of evidence indicating that phosphorylation of NRT1.1 at T101 affects NRT1.1-modulated LR development by altering NRT1.1 subcellular distribution, PM dynamics, and subcellular trafficking in LR cells to alter nitrate-dependent auxin transport and nitrate-regulated signaling.

LR plasticity plays a key role in enabling plants to obtain nutrients and survive in complicated surroundings (Stoeckle et al., 2018). Nitrate and auxin can have various functions in plant morphogenesis through their regulation of LR growth (Sun et al., 2017; Du and Scheres, 2018). Recently, genetic and physiological evidence has suggested that NRT1.1 represses LR growth by promoting basipetal auxin transport in LRs at LN (Krouk et al., 2010). The model was preliminarily explained by the repression of NRT1.1 protein accumulation in LR primordia (LRP) through posttranscriptional regulatory mechanisms by HN (Bouguyon et al., 2016). Furthermore, the point mutation of NRT1.1^T101A NRT1.1 was shown to inhibit its auxin transport facilitation, thereby leading to misregulation of DR5 activity in LRP and emergence of LR in absence of NO₃⁻ (Bouguyon et al., 2015). Here, we provided more in-depth understanding for the role of NRT1.1 phosphorylation in LR development through high-resolution NMT analysis and LR [³H]IAA uptake assay in combination with DR5-GFP fluorescence intensity quantification. We found that plants producing the nonphosphorylatable T101A mutant version of NRT1.1 exhibited increased LR density and auxin accumulation in LRP and new emerged LR resulting from reduced basipetal auxin transport in response to NO₃⁻ limitation (0 and 0.2 mM), whereas plants producing the phosphomimetic T101D mutant of NRT1.1 displayed cessation of LR elongation and auxin accumulation due to increased auxin transport (Fig. 1). Given the differences in LR growth between T101A and T101D plants, we conclude that the switch between the phosphorylation states of NRT1.1 present on the PM is also involved in the comprehensive model of how NRT1.1 acts to modulate LRP development.

An important determinant for controlling the fundamental properties of PM proteins is thought to be the spatiotemporal organization of these proteins. Bouguyon et al. (2015) showed that P492L mutation of NRT1.1 led to repression of LR development by preventing auxin transport, which might be due to its mistargeting from PM to the intracellular localization. Moreover, the mutation of P492L had no significant influences on primary nitrate response (PNR) and regulation of the genes of clusters 13 and 17 (Bouguyon et al., 2015), suggesting the possible role of NRT1.1 accumulation in intracellular compartments might partly contribute to its mediated NO₃⁻ signaling. These observations provide clues to indicate that the subcellular distribution is associated with the function of NRT1.1.

Protein phosphorylation is one of the most important and best-characterized posttranslational modifications. Of first importance, phosphorylation could induce the electrostatic and conformational changes of proteins. For NRT1.1, crystal structure studies reveal that T101 phosphorylation controls its dimerization and/or structural flexibility, thereby affecting the intrinsic NO₃⁻ transport capacity (Parker and Newstead, 2014; Sun et al., 2014). Phosphorylation of PM proteins also plays a vital role in a wide range of cellular processes including subcellular trafficking and PM dynamics (Offringa and Huang, 2013). Here, we showed that as the active form of NRT1.1 in response to LN and HN, respectively, phosphorylated and dephosphorylated, NRT1.1 exhibited the distinct behavior of LR development (Fig. 1). These data raises the question of whether the phosphorylation state of NRT1.1 could induce changes in its PM dynamics and subcellular trafficking, and specifically affect NO₃⁻-dependent auxin transport and NO₃⁻ signaling involved in LR development regulation. Thus, an intensive study by single molecular analysis was performed to provide strong support for our hypothesis.

Our protein labeling and SPT analysis data indicated that T101D clusters had fast lateral mobility with long motion range, high particle velocity, and reduced diffusion coefficient, which may enhance auxin transport efficiency. Conversely, T101A clusters had shorter lifetime and showed more stable behavior with limited motion range, low particle velocity, and increased diffusion coefficient, which are favorable for its dimerization and internalization (Fig. 2, A–E; Supplemental Fig. S6, A–C). The crystal structure of NRT1.1 indicates that under LN, nonphosphorylated NRT1.1 forms a homodimer with low structural flexibility, and phosphorylation at T101 may decouple the dimer and increase its flexibility (Tsay, 2014). Using subunit counting analysis, we found that both T101A and T101D proteins exhibited the mix of monomeric and dimeric status, and the percentage of dimer for T101A was significantly higher than that for T101D, suggesting that the phosphorylation state of NRT1.1 exerts significant effect on the interconversion between monomer and dimer state on the PM of LR cells (Fig. 2F; Supplemental Fig. S6F).

Membrane microdomains, sterol- and sphingolipid-enriched regions in the PM, can form platforms for clustering PM proteins and regulating the spatiotemporal dynamics of PM proteins (Bücherl et al., 2017). Here, we investigated the spatial relationship between phosphorylated and nonphosphorylated NRT1.1 and AtRem1.3, a commonly accepted marker of membrane microdomains (Demir et al., 2013). In quantitative colocalization assays, we found higher correlation coefficients for T101D-GFP with AtRem1.3-mCherry than for T101A-GFP with AtRem1.3-mCherry under both LN and HN (Fig. 3, A and B). Using FRET-FLIM, we found that the transgenic plants coexpressing T101D-GFP and AtRem1.3-mCherry had a shorter GFP lifetime than those coexpressing T101A-GFP and AtRem1.3-mCherry (Fig. 3, C and D). Considering previous reports that the most highly ordered lipid structures in cells of the transition zone correspond with high auxin flux (Mancuso et al., 2007; Zhao et al., 2015; Baluška et al., 2018), we speculate that phosphorylated NRT1.1 may aggregate into the PM microdomain in a process triggered by LN, which might provide an efficient means for transporting auxin.

Studies on PM dynamics of plant proteins and specifically on the role of phosphorylation in this process have only been initiated more recently. Xue et al. (2018) showed that the phosphorylation of blue light receptor phot1 enhanced its interaction with AtRem1.3 and promoted faster movement on the PM. The distinct behaviors of NRT1.1 phosphorylation variants, including dwell time, lateral diffusion, assembly state, and partitioning pattern were found in our study, indicating a nonnegligible role of phosphorylation on the PM spatiotemporal dynamics of NRT1.1 (Figs. 2–4). Given our finding of auxin transport activity difference between T101A and T101D, it is reasonable to propose that the spatiotemporal features of NRT1.1 on the PM are important for its auxin transport capacity.

Phosphorylation of PM proteins can regulate their sorting and subcellular localization, either indirectly by enhancing the exposure of sorting signals through conformational changes, or directly by enhancing the binding of regulatory proteins to the phosphorylated sorting signal itself (Cadena et al., 1994; Nesterov et al., 1995; Moeller et al., 2011). Phosphorylation might affect the endocytosis of PM proteins, either positively or negatively (Nguyen et al., 2013; Offringa and Huang, 2013). In this investigation, the T101D mutant showed a longer dwell time and constitutive endomembrane trafficking with slow recycling via a BFA-sensitive pathway, whereas T101A had a much shorter lifetime and underwent HN-induced internalization that was largely independent of ARF-GEFs (Fig. 4). These results provide support for the hypothesis that the nonphosphorylated form of T101 facilitates rapid internalization of NRT1.1, which may attenuate NRT1.1’s capacity for auxin transport.

A recent study indicates that both CME and CIE pathways are involved in PM protein internalization (Fan et al., 2015). In our study, we found that under LN conditions, almost 88.5% of T101D internalization was blocked in the chc2 mutant, implying that the CME pathway has a dominant role in the endocytosis of phosphorylated NRT1.1. After short-term HN treatment, mutation of CHC2 inhibited T101A endocytosis considerably (by ∼33.8%), suggesting that the CIE pathway plays an auxiliary, although not trivial, role in nonphosphorylated NRT1.1 internalization. We also found a high colocalization coefficient and cross correlation between T101A-GFP and AtFlot1-mCherry, implying that microdomain-associated endocytosis participates in the internalization of T101A as well (Fig. 5; Supplemental Fig. S9, A–D). After prolonged exposure to HN, T101D exhibited no visible vacuolar targeting, whereas T101A followed an endocytic trafficking route from the PM to the vacuole, passing mainly through the MVB, PVC, and LE and only partially via the TGN and EEs (Fig. 6), which is in good agreement with the results of other recent studies (Takano et al., 2005; Mbengue et al., 2016; Ortiz-Morea et al., 2016). Together, our data reveal a dual role of phosphorylation state in NRT1.1 endocytic pathways and intracellular trafficking routes.

Phosphorylation-mediated subcellular trafficking in plant cells was provided by studies on several functional proteins such as receptor kinase FLS2 and transporters PIP2;1 and PINs (Robatzek et al., 2006; Prak et al., 2008; Dhonukshe et al., 2010). From these events, specific phosphorylation has been shown to function in ligand or stress-induced endocytosis and apical recycling for polar targeting. Our findings provide new insights into phosphorylation-regulated PM protein trafficking in plants. We not only revealed the effects of phosphorylation on constitutive and HN-induced endocytosis and exocytosis of NRT1.1, but also on its endocytic pathways as well as intracellular trafficking routes (Figs. 5 and 6).

Receptor-mediated endocytosis is considered an integral part of signal transduction (Mukhopadhyay and Riezman, 2007; Huang and Chen, 2012; Irannejad et al., 2015; Paez Valencia et al., 2016; Dubeaux and Vert, 2017). However, the phosphorylation states and endocytic processes of NRT1.1 involved in nitrate-sensing systems have not been identified. Our results show that nonphosphorylated NRT1.1 promotes increases in [Ca²⁺]_cyt and ANR1 expression (Fig. 7), supporting the possibility that the T101 phosphorylation site of NRT1.1 has a role in fine-tuning nitrate-dependent signaling. In addition, we found that inhibiting T101A endocytosis decreased nitrate signaling, whereas increased vesicular accumulation of T101A increased nitrate signaling (Fig. 8, A–F), confirming that the endosome-localized NRT1.1 retains signaling activity before recycling or degradation. Notably, LR growth was greatly reduced in T101A-GFP/chc2-1 seedlings (Fig. 8, G and H), revealing that clathrin is required for PNR in response to LR development. Based on these results, we proposed a novel role of endocytosis in regulating nitrate signaling (modeled in Fig. 9): under LN, phosphorylated NRT1.1 proteins undergo slow endocytosis, which contributes to constitutive nitrate signaling, whereas under HN, the induced endocytosis of nonphosphorylated NRT1.1 proteins promote their entry into MVBs and/or LEs, which probably perpetuates PNR. With recent findings that intracellular localization of NRT1.1^P492L does not affect some downstream signaling, our results also provide evidence for the hypothesis that NRT1.1-mediated signaling partly initiates from endosomes rather than PM.

Figure 9.

Schematic model of phosphorylation-state–mediated NRT1.1 vesicle cycling and nitrate-dependent signal response for LR development. Blue solid arrows represent the direction of endocytosis and pink solid arrows represent the direction of secretion or recycling; Blue solid arrows over the PM represent more activity of NRT1.1 that moves down each branch through the indicated pathway. Arrowheads on the black lines represent activation and the vertical crossbars at the ends of lines represent inhibition. Dotted lines indicate attenuated responses in LN compared with HN. LN conditions (0.2 mm of NO₃^–). HN conditions (10 mm of NO₃^–).

Collectively, the results of this study identify a role for the different phosphorylation states of NRT1.1 in its PM spatiotemporal dynamics and subcellular trafficking in LR cells and suggest that this modulation mechanism is a novel strategy to control NRT1.1-regulated LR development. Under LN conditions, phosphorylation of T101 promotes NRT1.1 recruitment into functional membrane microdomains at the PM. These actions could facilitate NRT1.1-dependent auxin flux, thus depleting auxin levels in LRP and inhibiting their outgrowth. With an increase in nitrate, the nonphosphorylated NRT1.1 displays oligomerization and low lateral mobility at the PM, and undergoes a faster, inducible endocytosis. These behaviors could promote LR development by suppressing NRT1.1-auxin transport activity on the PM and stimulating Ca²⁺-ANR1 signaling from the endosome (Fig. 9).

MATERIALS AND METHODS

Plasmid Constructs

To create transgenic plants expressing NRT1.1, NRT1.1^T101D (T101D), and NRT1.1^T101A (T101A) fusions with GFP, the 1,751-bp coding region fragment was PCR-amplified from the wild-type NRT1.1 complementary DNA (cDNA) and the point mutations T101D and T101A were introduced using the recombinant PCR technique described in Vaucheret et al. (1990) and Ho et al. (2009). After that, the NRT1.1, T101D, and T101A cDNAs were each subcloned into a modified pCAMBIA 2300-GFP vector under the control of its native promoter region (∼3 kb) to generate the pNRT1.1::NRT1.1, pNRT1.1::T101D, and pNRT1.1::T101A constructs, respectively, and then introduced into the chl1-5 mutant plants by using Agrobacterium tumefaciens GV3101 to generate the transgenic plants expressing NRT1.1-GFP, T101D-GFP, and T101A-GFP (Supplemental Fig. S3A). For expression of NRT1.1, T101A, and T101D fusions with mRuby in transgenic yeast, NRT1.1, T101A, and T101D cDNA fragments amplified by PCR from the pNRT1.1::NRT1.1, pNRT1.1::T101A, and pNRT1.1::T101D-GFP constructs were subcloned into the modified binary vector pESC-mRuby-URA to create the pESC-NRT1.1-mRuby-URA, pESC-T101A-mRuby-URA, and pESC-T101D-mRuby-URA vectors, respectively, which were then transferred into Saccharomyces cerevisiae strain YPH 499 (Supplemental Fig. S1B).

Transgenic Lines

The wild-type Arabidopsis (Arabidopsis thaliana) accession used was Columbia (Col-0). The transgenic Arabidopsis mutant lines NRT1.1^T101A (T101A) and NRT1.1^T101D (T101D) and the NRT1.1 total deletion mutant chl1-5 were kindly provided by Yi-Fang Tasy (Ho et al., 2009; Supplemental Fig. S1A). The transgenic line DR5::GFP was kindly provided by Rujin Chen (Laxmi et al., 2008). Transgenic lines expressing DR5::GFP were obtained by crossing the DR5::GFP line with the chl1-5, T101A, or T101D lines. The chc2-1 mutant seeds were provided by Jiří Friml (Ghent University; Kitakura et al., 2011). T101D-GFP/chc2-1 and T101A/chc2-1 were obtained by crossing the T101A-GFP and T101D-GFP lines with chc2-1. The fluorescent reporters AtRem1.3-mCherry and Flot1-mCherry were developed in our laboratory (Wang et al., 2015; Lv et al., 2017). Lines expressing TGN, EE, MVB, LE, PVC, and vacuole markers were VTI12-mRFP (Zouhar et al., 2009), VHA-a1-mRFP (Dettmer et al., 2006), CLC-mCherry (Wang et al., 2015), AtSNX1-mRFP (Jaillais et al., 2008), mRFP-SYP22 (Ebine et al., 2008), and RabG3f-mRFP (Geldner et al., 2009), respectively. For dual-color imaging, transgenic Arabidopsis plant carrying T101A-GFP were crossed with AtRem1.3-mCherry, Flot1-mCherry, VTI12-mRFP, VHA-a1-mRFP, CLC-mCherry, AtSNX1-mRFP, mRFP-SYP22, and RabG3f-mRFP plants. The Arabidopsis transgenic line ATG8-CFP (expressing a fusion of the autophagosome marker ATG8 and cyan fluorescent protein [CFP]) was kindly provided by Yule Liu (Wang et al., 2013b). Lines expressing NRT1.1-GFP and ATG8-CFP were obtained by crossing the ATG8-CFP line with the NRT1.1-GFP line.

Plant Growth Conditions

Arabidopsis seeds were routinely surface-sterilized for 3 min in 4:1 (v/v) 85% ethanol:H₂O₂, plated on basic medium containing 12.5 mm of (NH₄)₂ succinate, 0.5 mm of CaSO₄, 0.5 mm of MgCl₂, 1 mm of KH₂PO₄, 2.5 mm of MES at pH 6.5, 50 μm of NaFeEDTA, 50 μm of H₃BO₃, 12 μm of MnCl₂, 1 μm of CuCl₂, 1 μm of ZnCl₂, and 0.03 μm of NH₄MoO₄, then chilled at 4°C for 2 d and transferred to a growth room with a photoperiod of 16 h (light, 100 μmol m⁻² s⁻¹)/8 h (dark) and temperature of 23°C/20°C. After the indicated time of growth (depending on the experiment), the seedlings were shifted to a version of the same growth medium in which the (NH₄)₂ succinate was replaced with different concentrations of KNO₃ (0, 0.2, 1, and 10 mm; Ho et al., 2009).

IAA Uptake Assay in Yeast

The transformed yeast cells were grown to log phase in synthetic dextrose dropout media without uracil (SD-URA) and then transferred to synthetic galactose dropout media without uracil (SG-URA; SD-URA containing 2% [m/v] Gal instead of dextrose) for 24–48 h to induce NRT1.1, T101A, T101D, and mRuby expression, and then harvested by centrifugation and resuspended in SG-URA. For each assay, 10 μL of resuspended cells were immobilized on a coverslip for 5 min, washed off with standard medium (0.5 μm of IAA, 2% [m/v] Gal, 0.3 mm of MES, at pH 5.8) to leave a monolayer of attached yeast cells, and then incubated in the standard medium for 10 min. The kinetics of net IAA flux of each cell population were measured by NMT (Beijing Science and Technology) and analyzed with the program “Mage Flux” (http://www.xuyue.net/mageflux). The experiment was replicated six times for each category of transformed cell.

IAA Uptake Assay in LRs

Wild-type, T101A, and T101D seedlings were grown on basic medium containing different concentrations of KNO₃ (0.2 and 1 mM) for 10 d. Agar blocks of 1 mm in diameter supplemented with 100 nm of tritium-labeled IAA ([³H]IAA, cat. no. NET1175; Perkin-Elmer) were placed next to the LR tips (Supplemental Fig. S1D). After incubation for 1.5 h in the dark, agar blocks were removed. Meanwhile, 0.3 mm of each LR tip was excised and discarded, and then a 1-cm root segment was excised from the remaining LR and placed into a vial containing 2 mL of scintillation fluid. The radioactivity level of the root segment was measured with a model no. 1450 MicroBeta Liquid Scintillation Counter (Perkin-Elmer) for 1 min. The experiment was repeated six times for each category of plant. Six seedlings for each genotype and treatment were detected and presented.

Confocal Microscopy

Confocal microscopy was done with a model no. TCS SP8 Confocal Microscope (Leica, Germany) fitted with 63× and 100× oil-immersion objectives. A fluorescence recovery after photobleaching assay of the GFP-labeled seedlings was performed using a laser (488 nm) for excitation and detecting light emission at 505–545 nm. The region of interest (ROI) was selected and bleached for 15 s with a 488-nm laser at 100% laser power after a prebleach of 3 s. A 2-s time interval was used for monitoring fluorescence recovery. The fluorescence recovery was quantified using ImageJ (1.51w; Collins, 2007). OriginPro v8.5 was used for curve fitting (Stevenson, 2011). For the mRuby fluorescence assay in yeast, mRuby was excited using the 563-nm line of an argon laser and detected from a 581-nm bandpass filter (red). For GFP fusions and Fluo-4-AM-dyed seedlings, an argon laser (488-nm line) was used for excitation and the emission light from 505 to 545 nm (green) was detected with a 63× objective (HCX PL APO 1.4 NA; Leica). Confocal imaging of Arabidopsis LR cells expressing GFP with FM4-64 dye treatment was performed with excitation at 488 nm. The fluorescence emissions were detected with the spectral detector set BP 505–545 (green) and LP 560–640 (red; Leica). Confocal imaging of Arabidopsis LR cells expressing GFP with mCherry or red fluorescent protein (mRFP) was performed via excitation with 488- and 585-nm light, respectively (multitrack mode) and detection of fluorescence emissions at 505–545 nm (GFP) and 600–630 nm (mCherry or mRFP; Leica). Arabidopsis LR cells coexpressing ATG8-CFP and NRT1.1-GFP were excited with 436-nm and 488-nm light and detected with 450–476 (CFP) and 505–545 (GFP).

Fluo-4 am-Based Ca²⁺ Imaging in LRs

For [Ca²⁺]_cyt signal imaging of the LR cells of wild-type, chl1-5, T101A, and T101D seedlings, seedlings were grown on the normal half-strength Murashige and Skoog (MS) medium for 8 d, transferred to liquid basic medium without nitrate for 2 d, then incubated with Fluo-4 am for 20 h at 4°C in the dark. After three repeats of a wash-out with basic medium followed by 1 h of standing, each seedling was plated on a slide and covered with a cover glass, and then basal medium (200 μL) with 0, 0.2, and 10 mm of KNO₃ was loaded along one edge of the coverslip. A piece of absorbent paper was placed at the opposite edge to wick out the buffer. For [Ca²⁺]_cyt signal imaging of the effects of inhibitor treatment, the inhibitor was added for 1 h and then the plants were treated with 10 mm of KNO₃ before imaging. Confocal imaging was acquired using a model no. SP8 Confocal Microscope (Leica). The fluorescence intensity was determined with the “ROI” function of ImageJ (1.51w) for each seedling. The intensity data were exported and processed using the program OriginPro v8.5. The modified rainbow RGB Lookup Table was applied with ImageJ. The images were exported and processed using the software Adobe Illustrator (CC 2017).

RNA Isolation and RT-qPCR

For ANR1 expression in LRs, wild-type, chl1-5, T101A, and T101D seedlings were grown in basic medium at pH 5.8 for 10 d, transferred into fresh liquid basic medium at pH 5.5 overnight/16 h and then placed for 15 min in the same growth medium but with the (NH₄)₂ succinate replaced with the appropriate concentration of KNO₃ (depending on the experiment). The apical parts (10–15 mm) of the plant LRs were separated surgically from the rest of the root system (Remans et al., 2006). For NRT1.1 and ANR1 expression in GFP-labeled transgenic plants, 7-d–old plants grown on basic medium were harvested and then transferred to liquid medium of appropriate composition. Total RNA was extracted from homogenized tissues by using the RNA prep Pure Plant Kit (Tiangen Biotech). Three micrograms of total RNA were used to prepare cDNA using a PrimeScript first-Strand cDNA Synthesis Kit (Takara Biomedical Technology) by reverse transcription with Moloney-Murine Leukemia Virus reverse transcriptase and oligo(dT)₁₈ primers. Gene expression was determined by PCR and quantitative real-time PCR with a CFX Connect Real-Time PCR system (Bio-Rad) by SYBR Premix Ex Taq II (Tli RNase H Plus; Takara Biomedical Technology). ACTIN2, CLATHRIN, and UBIQUITIN10 served as the controls for RT-qPCR and gene-specific primers are listed in Supplemental Table S1. The experiments were repeated six times, and the 2^–ΔΔCT quantification method was used to evaluate quantitative variation.

Analysis of LR Growth

Seedlings were germinated on half-strength MS medium for 3 d and then transferred to basic medium containing different concentrations of KNO₃ (0, 0.2 and 1 mm) for an extra 5 d. The root systems were scanned at 500 dpi (model no. EOS 600D; Canon). Root growth parameters including LR length and density were analyzed using the software ImageJ. Fifteen seedlings for each genotype and treatment were measured and presented. The experiment was repeated seven times.

VA-TIRFM and Fluorescence Image Analysis

VA-TIRFM was performed on an inverted microscope (model no. IX-71; Olympus) with a total internal reflective fluorescence illuminator (model no. IX2-RFAEVA-2; Olympus) and a 100× oil-immersion objective (numerical aperture = 1.45; Olympus). GFP-labeled proteins in LR epidermal cell from 12-d–old Arabidopsis seedlings were excited with 488-nm laser lines from a diode laser (Chang-chun New Industries Optoelectronics Technology) and their emission fluorescence was obtained with a BA510IF (505/50) filter for GFP. The fluorescence signals were detected with a back-illuminated EMCCD camera (model no. iXon DV8897D–CS0–VP; Andor Technology). Images were acquired with 100-ms exposure times and a 330-ms time-lapse series of single particles of T101D-GFP or T101A-GFP, which was taken with up to 100 images per sequence. For the analysis of VA-TIRFM images, the background was subtracted with a “rolling-ball radius: 50 pixels,” and pseudo-colors (green) were added. The stand-alone MATLAB Graphical User Interface program (R2014a; https://neurophotonics.ca/software) was then used for SPT according to the method described by Jaqaman et al. (2008). Kymograph and lifetime analyses were performed as described in Eichel et al. (2016). The diffusion coefficient, motion range and particle velocity analyses were done according to methods described in Li et al. (2011), Hao et al. (2014) and Lv et al. (2017). The single-particle fluorescence intensity and the photobleaching steps were calculated according to the methods described in Wang et al. (2015) and Lv et al. (2017). The Progressive Idealization and Filtering program was used for step-wise photobleaching analysis after background subtraction as described in McGuire et al. (2012).

FCCS Analysis

To quantitatively measure the interactions among T101A-GFP and AtCLC-mCherry, AtFlot1-mCherry, and AtRem1.3-mCherry, LR cells of 10-d–old seedlings expressing T101A-GFP with AtCLC-mCherry/AtFlot1-mCherry/AtRem1.3-mCherry were detected by FCCS, performed on a model no. TCS SP5 microscope (Leica) as described in Mütze et al. (2011). GFP and mCherry were excited with 488-nm and 561-nm laser lines, respectively. Emission filters were set as BP 505–540 and LP 580 for the green and red channels, respectively. The relative cross correlation ([Gc(0)]/[Gg(0)]) was calculated by normalizing the amplitude of the cross-correlation function to the amplitude of the autocorrelation function of GFP as described in Li et al. (2011).

Transient Infiltration of Nicotiana benthamiana Leaves

Leaves of 7-week–old N. benthamiana were infiltrated with equal amounts of A. tumefaciens expressing full-length NRT1.1-GFP, T101A-GFP, or T101D-GFP fusion constructs together with AtRem1.3-mCherry, AtFlot1-mCherry, or AtCLC-mCherry with all the possible combinations. After 2 d of inoculation, leaves were harvested and treated with 0.2- and 10-mm nitrate and then photographed using a model no. SP8 Confocal Microscope (Leica).

FRET-FLIM

For FRET-FLIM, seedlings coexpressing AtRem1.3-mCherry and T101A-GFP or T101D-GFP were treated with the appropriate concentration of nitrate. FLIM was performed on an inverted model no. FV1200 microscope (Olympus) equipped with a model no. picoHarp300 controller (PicoQuant). The excitation at 488 nm was carried out by a picosecond pulsed diode laser at a repetition rate of 40 MHz through a water-immersion objective (60×, numerical aperture = 1.2). The emitted light was filtered with a 510/55-nm bandpass filter and detected with an Micro Photon Devices Single Photon Avalanche Diodes detector. Images of each selected ROI were obtained with acquisition of at least 50,000 photons. From the fluorescence intensity images, the decay curves were calculated per pixel and fitted with either a mono- or double-exponential decay model using the software “SymphoTime 64” (PicoQuant). The monoexponential model function was applied for donor samples with only GFP present and the double-exponential model function was used for samples containing GFP and mCherry. Data on fluorescence lifetime (in picoseconds) were obtained and FRET efficiency (%) analysis was performed using the software Excel 2013 (Microsoft).

Colocalization Analysis

Colocalization analysis of confocal images was performed with the software ImageJ. For transient infiltration of N. benthamiana leaves, NRT1.1-GFP, T101A-GFP, and T101D-GFP fluorescence colocalized with AtRem1.3-mCherry were calculated by using the plug-in “Coloc 2” (https://imagej.net/Coloc_2). The obtained modified Manders coefficients were interpreted as the fraction of colocalization for both channels (i.e. colocalization of AtRem1.3-mCherry with NRT1.1-GFP, T101A-GFP, or T101D-GFP and vice versa; Bücherl et al., 2013). Pearson correlation coefficients were obtained as another measure of colocalization (Bücherl et al., 2013, 2017; Demir et al., 2013). The colocalization analysis for T101A-GFP and AtFlot1-mCherry, AtCLC-mCherry, and AtRem1.3-mCherry in LR cells was carried out as described using ImageJ in Jarsch et al. (2014) and Bücherl et al. (2017) . Endomembrane compartments were excluded from the colocalization analysis by ROI selection. First, acquired images were “Mean-Shift” filtered with a radius of 2 pixels. Background was subtracted using the “Rolling-ball” method with a radius of 20 pixels. ROIs were manually selected and the colocalization plug-in “Intensity Correlation Analysis” was applied for quantification (Bücherl et al., 2017; the plug-in is available at: http://www.aomf.ca/WCIF.html). For the colocalization analysis between T101A-GFP with different endomembrane markers in LR cells, the plug-in “PSC colocalization” in the software ImageJ was used to obtain the Pearson correlation coefficients as colocalization readouts. To determine the percentage of T101A-GFP vesicles colocalized with different endomembrane markers, each individual T101A-GFP spot was selected as an individual ROI and colocalization was analyzed with a threshold of 10, considering the spots for which the Pearson correlation value was >0.5 as positively labeled endosomes. A threshold of 10 was used to avoid noise, and calculations were done by measuring the background gray values in the analyzed images.

Drug Treatments

BFA and CHX were obtained from Sigma-Aldrich and dissolved in 100% DMSO to yield 50-mm stock solutions that were diluted to 50 μm with half-strength liquid MS medium for use. A solution of 200 mm of MβCD (Sigma-Aldrich) was prepared in deionized water and then diluted to 10 mm with half-strength liquid MS medium for use. FM4-64 (Invitrogen) was kept as a 5-mm stock solution and diluted to a 5-μm working solution with half-strength liquid MS medium. Inhibitors were used at the following concentrations: 2 μm of Conc A (Sigma-Aldrich; with 2 mm of DMSO stock), 33 μm of WM (Sigma-Aldrich; with 33 mm of DMSO stock), 1 μm of Baf A1 (Tocris Biosciences; with 1 mm of DMSO stock), and 20 μm of MG132 (Sigma-Aldrich; with 20 mm DMSO stock).

Immunoblot Analysis

For immunoblot analyses, total proteins were extracted from LRs of 12-d–old seedlings (100–200 mg). For protein detection, the following antibodies (Abbkine Scientific) were used: anti-GFP tag rabbit polyclonal antibody (1/5,000) and antiplant actin mouse monoclonal antibody (1/10,000) as primary antibody, and IPKine HRP Goat Anti-Rabbit IgG HCS (1/10,000) and IPKine HRP Goat Anti-Mouse IgG HCS (1/10,000) as second antibody. The intensity of each band was detected with ImageJ and normalized with respect to the control.

Accession Numbers

Sequence data from this work can be found in The Arabidopsis Information Resource database under the following accession numbers: AT1G12110 (NRT1.1), AT2G14210 (ANR1), AT3G18780 (ACTIN 2), AT4G24550 (CLATHRIN), and AT4G05320.2 (UBIQUITIN 10).

Supplemental Data

The following materials are available.

• Supplemental Figure S1. T101D mutation strengthens the auxin transport activity of NRT1.1 in LRs.

• Supplemental Figure S2. Confocal images and quantitative analysis of DR5::GFP in LRs.

• Supplemental Figure S3. The roots of transgenic NRT1.1-GFP, NRT1.1^T101D-GFP, and NRT1.1^T101A-GFP plants have normal biological function and organizational structure.

• Supplemental Figure S4. The transgenic NRT1.1-GFP, NRT1.1^T101D-GFP, and NRT1.1^T101A-GFP plants have corresponding LR phenotypes.

• Supplemental Figure S5. Dynamics of T101A-GFP and T101D-GFP on the PM.

• Supplemental Figure S6. Motion parameters of T101A-GFP and T101D-GFP on the pm.

• Supplemental Figure S7. The phosphorylation state of NRT1.1 affects its colocalization with AtRem1.3.

• Supplemental Figure S8. Secretion and internalization of NRT1.1 are affected by its phosphorylation state.

• Supplemental Figure S9. Both CME and CIE pathways are involved in the endocytosis of T101D-GFP and T101A-GFP.

• Supplemental Figure S10. The degradation of NRT1.1 is involved in proteasome system and autophagy under HN conditions and nitrogen starvation, respectively.

• Supplemental Figure S11. Ca²⁺ signaling disappears within 120 s after nitrate treatment in LR epidermis cells of wild-type Arabidopsis.

• Supplemental Table S1. Primer used in this research.

• Supplemental Video S1. The movie of dynamics of T101D-GFP spots (green) on the PM under the treatment of 0.2-mm NO₃⁻ concentration.

• Supplemental Video S2. The movie of dynamics of T101A-GFP spots (green) on the PM under the treatment of 10-mm NO₃⁻ concentration.