# The Arabidopsis NRT1.1 transceptor coordinately controls auxin biosynthesis and transport to regulate root branching in response to nitrate

Amel Maghiaoui, Eléonore Bouguyon, Candela Cuesta, Francine Perrine-Walker, Carine Alcon, Gabriel Krouk, Eva Benkova, Philippe Nacry, Alain Gojon, Lien L. Bach

<figure><img src="image_1.png" /></figure>

## To cite this version:

Amel Maghiaoui, Eléonore Bouguyon, Candela Cuesta, Francine Perrine-Walker, Carine Alcon, et al.. The Arabidopsis NRT1.1 transceptor coordinately controls auxin biosynthesis and transport to regulate root branching in response to nitrate. Journal of Experimental Botany, 2020, 71 (15), pp.4480-4494. 10.1093/jxb/eraa242/5840670. hal-02619371

HAL Id: hal-02619371

https://hal.inrae.fr/hal-02619371

Submitted on 25 May 2020

HAL is a multi-disciplinary open access archive for the deposit and dissemination of scientific research documents, whether they are published or not. The documents may come from teaching and research institutions in France or abroad, or from public or private research centers.

L'archive ouverte pluridisciplinaire HAL, est destinée au dépôt et à la diffusion de documents scientifiques de niveau recherche, publiés ou non, émanant des établissements d'enseignement et de recherche français ou étrangers, des laboratoires publics ou privés.
The *Arabidopsis* NRT1.1 transceptor coordinately controls auxin biosynthesis and transport to regulate root branching in response to nitrate

Amel Maghiaoui*¹, Eléonore Bouguyon*¹,³, Candela Cuesta²,⁴, Francine Perrine-Walker¹,⁵,
Carine Alcon¹, Gabriel Krouk¹, Eva Benkova², Philippe Nacry¹, Alain Gojon¹ and Liên
Bach¹,**

* These authors contributed equally to this work.

¹BPMP, Univ Montpellier, CNRS, INRAE, Institut Agro, Montpellier, France

²Institute of Science and Technology Austria, Klosterneuburg, 3400, Austria

³Present address: Site de Bellepierre, UFR Santé, Université de la Réunion, 1 Allée des Aigues Marines, 97487 Saint Denis Cedex, France

⁴Present address: Instituto Universitario de Biotecnología de Asturias, Departamento de Biología de Organismos y Sistemas, Universidad de Oviedo, Spain

⁵Present address: School of Life and Environmental Sciences, F22 Life, Earth and Environmental Sciences Building, University of Sydney NSW 2006, Australia

** Address Correspondence to lien.bach@supagro.fr

The author responsible for distribution of materials integral to the findings presented in this
article in accordance with the policy described in the Instructions for Authors is: Liên BACH
(lien.bach@supagro.fr).
HIGHLIGHT

The NRT1.1 transceptor controls both upstream auxin biosynthesis for lateral root primordia emergence and downstream action of auxin for lowering the mechanical resistance of the primary root tissues allowing LRP outgrowth.
ABSTRACT

In agricultural systems, nitrate is the main source of nitrogen available for plants. Besides its role as a nutrient, nitrate has been shown to act as a signal molecule for plant growth, development and stress responses. In *Arabidopsis*, the NRT1.1 nitrate transceptor represses lateral root (LR) development at low nitrate availability by promoting auxin basipetal transport out of the LR primordia (LRPs). In addition, our present study shows that NRT1.1 acts as a negative regulator of the *TAR2* auxin biosynthetic gene expression in the root stele. This is expected to repress local auxin biosynthesis and thus to reduce acropetal auxin supply to the LRPs. Moreover, NRT1.1 also negatively affects expression of the LAX3 auxin influx carrier, thus preventing cell wall remodeling required for overlying tissues separation during LRP emergence. Both NRT1.1-mediated repression of *TAR2* and LAX3 are suppressed at high nitrate availability, resulting in the nitrate induction of *TAR2* and LAX3 expression that is required for optimal stimulation of LR development by nitrate. Altogether, our results indicate that the NRT1.1 transceptor coordinately controls several crucial auxin-associated processes required for LRP development, and as a consequence that NRT1.1 plays a much more integrated role than previously anticipated in regulating the nitrate response of root system architecture.

Key words: auxin signaling, development, lateral root primordia, nitrate, NRT1.1, transceptor
# INTRODUCTION

Nitrogen (N) is an essential element that plants can acquire from various inorganic or organic sources in the soil, but nitrate ($NO_3^-$) is the main form taken up and assimilated by most species (Nacry et al., 2013). Soil $NO_3^-$ concentration is often a major factor limiting the growth and yield of crop plants as it fluctuates dramatically in both time and space (Crawford and Glass 1998; Miller et al., 2007). To cope with this, plants are able to quickly modulate their $NO_3^-$ acquisition efficiency according to changes both in their own internal N status and in the external $NO_3^-$ availability (Forde 2002; Nacry et al., 2013; Kiba and Krapp 2016; O'Brien et al., 2016). In addition to the regulation of root $NO_3^-$ transport systems (Kiba and Krapp 2016; Bellegarde et al., 2017; Jacquot et al., 2017), modification of root system architecture (RSA) is another key component of this response (Nacry et al., 2013; Forde, 2014; Giehl and von Wirén, 2014; Kiba and Krapp, 2016; O'Brien et al., 2016). In more detail, $NO_3^-$ has a dual effect on RSA in many species. On the one hand, high $NO_3^-$ provision to the plant has a systemic repressive effect (Zhang and Forde, 1998; Giehl and von Wirén, 2014), and as a consequence mild $NO_3^-$ limitation results in a general stimulation of root growth for improved foraging of the soil (Forde, 2002; Hermans et al., 2006; Giehl and von Wirén, 2014). On the other hand, $NO_3^-$ supply locally stimulates growth of the roots in contact with a high $NO_3^-$ concentration, therefore contributing to preferential colonization and exploitation of the $NO_3^-$-rich areas of the soil (Drew, 1975; Forde, 2002; Ruffel et al., 2011, Nacry et al., 2013). In dicotyledonous species where the root system is generally constituted of a primary root (PR) originating from the embryo, onto which lateral roots (LRs) develop post-embryonically, $NO_3^-$ was generally reported to markedly affect LR development and growth, and to a much lesser extent PR growth (Forde, 2014). Furthermore, many responses of RSA to changes in $NO_3^-$ availability were shown to be due to a signaling rather than a nutritional effect, pointing out the crucial role of specific $NO_3^-$ sensing and signaling systems (O'Brien et al., 2016; Guan, 2017; Sun et al., 2017).

Compelling evidence indicates that the membrane $NO_3^-$ transporter NRT1.1 (CHL1/NPF6.3), first identified as an influx carrier participating in the root uptake of $NO_3^-$ (Tsay et al., 1993; Liu and Tsay, 2003), is a central component of the $NO_3^-$ sensing and signaling systems in *Arabidopsis thaliana* (Gojon et al., 2009, 2011; Nacry et al., 2013; Forde, 2014; Bouguyon et al., 2015). This led to the proposal that it acts as a $NO_3^-$ transceptor (transporter/receptor), with a dual transport/sensing function (Muños et al., 2004; Walch-Liu and Forde, 2008; Ho et
al., 2009; Wang et al., 2009; Gojon et al., 2011). In particular, NRT1.1 is a master player in the NO₃⁻ regulation of RSA as it governs many responses of LR growth to NO₃⁻ (Remans et al., 2006; Krouk et al., 2010; Bouguyon et al., 2015; O'Brien et al., 2016; Sun et al., 2017). In *Arabidopsis*, LRs originate from pericycle founder cells that divide and differenciate in order to initiate a LRP (Stage I), which has to develop across overlying tissues to eventually emerge out of the parent root (Stage VIII) (Malamy and Benfey, 1997). Through this series of well-defined stages, auxin plays a central role in controlling many steps of LRP development (Dubrovsky et al., 2008; Benkova et al., 2003; Bhalerao, 2002; Fukaki et al., 2007; Ljung et al., 2001). Within the LRP, an auxin gradient is progressively established with a maximum at the apex that is required for proper growth of the LRP (Benkova et al., 2003). This gradient is created by the coordinated activity of efflux proteins (encoded by the *PIN* and *ABCB* gene families) that facilitate movement out of cells and influx transporters (encoded mostly by the *AUX/LAX* gene family) that facilitate auxin entry into cells (Petrasek and Friml, 2009; Vieten et al., 2007). A “fountain model” was proposed for auxin flow in LRP based on auxin accumulation and auxin transporters expression patterns (Benkova et al., 2003). According to this model, an acropetal flow (involving PIN1/7 located in the inner cell layers) allows auxin to come from the PR stele and to reach the LRP apex where it accumulates, while a basipetal flow (mediated by PIN3/4 in the outer cell layer) redirects the hormone to the PR. In addition, AUX1, ABCB transporters and LAX3 have been shown to be involved in LRP development: AUX1 facilitates LRP initiation and emergence (Marchant et al., 2002; Swarup and Péret, 2012) and ABCB1 and ABCB19 physically interact with PIN1 as well with AUX1 for normal LR initiation and development (Mravec et al., 2008). The influx/efflux auxin transporter ABCB4 (Yang and Murphy, 2009; Wu et al., 2010; Kubes et al., 2012) was also identified as a negative regulator of LRP initiation (Santelia et al., 2005) through a mechanism remaining to be elucidated. Finally, LAX3 plays an important role in LR emergence process by increasing the level of auxin in LRP overlying cells, thus leading to a remodeling of cortical/epidermal cell walls that facilitates their separation and allows the outgrowth of the LRP (Swarup et al., 2008).

Within this general scheme, we recently proposed a model for the mechanism by which NRT1.1 regulates LR growth in response to NO₃⁻ (Krouk et al., 2010; Bouguyon et al., 2015; Bouguyon et al., 2016). Using heterologous expression systems, we found that NRT1.1 is not only able to transport NO₃⁻ but also auxin. The auxin influx activity of NRT1.1 is however inhibited by NO₃⁻ in a concentration-dependent manner (Krouk et al., 2010). Furthermore, we
observed that NRT1.1 is expressed in LRPs only in the absence or at low NO₃⁻ availability (Bouguyon et al., 2016). Taking into account the perfect match between the localization of the NRT1.1 protein with that of the basipetal transport route for auxin (outer cell layer of LRP), we postulated from the above observations that NRT1.1 mediates auxin influx into the outer cell layer of the LRPs, and thus facilitates the basipetal export of auxin out of these organs in the absence of NO₃⁻ or at low NO₃⁻ concentration (Krouk et al. 2010). Accordingly, in the absence of NO₃⁻ or at low NO₃⁻ concentration NRT1.1 lowers auxin accumulation/signaling (assessed by DR5 activity) in the LRPs, which in turn leads to an inhibition of the emergence of the LRPs (Krouk et al., 2010; Bouguyon et al., 2015; Bouguyon et al., 2016; Zhang et al., 2019). At higher (>0.5 mM) NO₃⁻ supply, both NRT1.1 expression in LRPs and NRT1.1-dependent auxin transport activity are inhibited. This results in an increased auxin accumulation/signaling in LRPs, which favors their emergence. Therefore, the local stimulation of LR growth by NO₃⁻ is explained by the relief of the repressive action of NRT1.1. Accordingly, null mutants for NRT1.1 (chl1 mutants) have a much higher LR density than wild-type plants in the absence, but not in the presence of NO₃⁻ (Krouk et al. 2010, Bouguyon et al. 2015, Bouguyon et al. 2016).

An important point that remains unclear in this model is to know if, in addition to its direct effect on auxin transport in the LRPs, NRT1.1 also acts indirectly through modified expression/activity of other molecular actors determining auxin distribution in these organs. As matter of fact, auxin transport in LRP also involves the action of up to ten other auxin carriers, i.e., PIN1-4, PIN7, ABCB1, ABCB4, ABCB19, AUX1 and LAX3 (see Petrášek and Friml, 2009). Interestingly, earlier transcriptome studies at the whole root or whole plant level have revealed that several auxin transporter genes are responsive to NO₃⁻ (Gutierrez et al., 2007) in an NRT1.1-dependent manner (c Furthermore, there are reports indicating that responses of RSA to nutrients, including N, can be accounted for by changes in expression of AUX and PIN genes (Li et al., 2011; Giehl et al., 2012; Sun et al., 2017; Naz et al., 2019). Nevertheless, these considerations do not allow any conclusion on a putative role of NRT1.1 in regulating other auxin carriers for modulating LR development because little is known on the regulation of PIN, ABCB and AUX/LAX expression by NO₃⁻ signaling in LRPs. In addition, recent evidence highlights the importance of not only auxin transport, but also local auxin biosynthesis in determining the changes in auxin accumulation in plant tissues in response to external factors (Brumos et al., 2018; Zhao et al., 2018). This suggests that the nutrient regulation of RSA may also rely on a control exerted on auxin biosynthesis pathways. Of particular interest is the observation that the *Arabidopsis* TAR2 gene, encoding
a Tryptophane amino transferase catalyzing the first step of the Trp-dependent auxin biosynthetic pathway, is required for stimulation of LR growth in response to mild N limitation (Ma *et al.*, 2014). However, nothing is known on the role of this gene in the local $NO_3^-$ stimulation of LRP development in connection with NRT1.1-dependent signaling.

In the current work, we thus aimed at investigating how $NO_3^-$ regulates the expression of the major auxin transporters (PINs, ABCBs, AUX1/LAX3) and of the TAR2 auxin biosynthetic enzyme during LRP development, and at determining if this regulation may account for an indirect effect of NRT1.1 on LR growth, through its $NO_3^-$ signaling action on auxin transporter or auxin biosynthetic enzyme expression. As shown by the results detailed below, our work revealed that NRT1.1 has a much more integrated role in regulating LRP development than anticipated before. Indeed, our data indicate that NRT1.1 not only participates in auxin transport in LRPs, but also controls both the upstream biosynthesis of the hormone as a source of auxin for the LRPs, and the downstream action of auxin for lowering the mechanical resistance of the PR tissues to LRP outgrowth.

## MATERIALS & METHODS

### Plant material

The *Arabidopsis thaliana* accessions used in this study were Columbia (Col-0) and Wassilewskija (WS). The following GFP-tagged lines were used and crossed with the *chl1-5* mutant (Tsay *et al.*, 1993): pDR5::VENUS (Brunoud *et al.*, 2012), pPIN1::PIN1-GFP (Benkova, 2003), pPIN2::PIN2-GFP (Xu and Scheres, 2005), pPIN3::PIN3-GFP (Zadnikova *et al.*, 2010), pPIN4::PIN4-GFP (Vieten, 2005), pPIN7::PIN7-GFP (Blilou *et al.*, 2005; Mravec *et al.*, 2008), pAUX1::AUX1-YFP (Swarup, 2004), pLAX3::LAX3-YFP (Swarup *et al.*, 2008), DR5::GFP (Ulmasov *et al.*, 1997), pTAR2::GUS and tar2-c (Ma *et al.*, 2014). The following GFP-tagged lines were used and crossed with the *chl1-10* mutant (Munos *et al.*, 2004): pABCB1::ABCB1-GFP (Mravec *et al.*, 2008), pABCB4::ABCB4-GFP (Cho *et al.*, 2007), pABCB19::ABCB19-GFP (Bailly *et al.*, 2008). Homozygous plants for both *chl1-5* mutation and genes encoding tagged proteins were screened on F2 and F3 offsprings by PCR for *chl1-5* deletion as described in Mounier *et al.*, (2014) and fluorescence. Transgenic Arabidopsis line expressing *NRT1.1-GFPLoop* in WT background were obtained as described by Bouguyon *et al.*, 2016.
## Plant culture and root growth analysis

Seedlings were grown for 12 days in vertical petri dishes according to Krouk *et al.*, (2010). Basal culture medium without N was supplemented with the appropriate concentration of KNO3, or Gln for each experiment as described in figure's caption. For analysis of root system architecture, the petri dishes were scanned using a Epson Perfection V700 PHOTO scanner (24-bits-color; resolution 600 ppp). The root system parameters such as primary root length, lateral roots length and number as well as lateral root density were measured by the Neuron J plugging from Image J software. Root system pictures for all genotypes were taken using Nikon digital camera D90. To analyze root growth parameters, Student T Test's was used to show statistical difference between genotypes and treatments.

## RNA analysis

Root samples were frozen in liquid nitrogen and disrupted for 1 min at 30 s⁻¹ in a Retch mixer mill MM301 homogenizer. Total RNA was extracted using TRIzol reagent (Invitrogen), DNase treated (Qiagen) purified using an RNeasy MinElute Cleanup Kit (Qiagen) and reverse transcription was achieved with 4 µg of RNAs with ThermoScript™ RT-PCR System for First-Strand cDNA Synthesis (Invitrogen) using an anchored oligo-dT(20) primer. Accumulation of transcript was measured by quantitative real-time PCR (LightCycler 480; Roche Diagnostics) using the Light Cycler 480 SYBR Green 1 Master kit (Roche Diagnostics). All steps were performed according manufacturer's recommendations. The gene expression was normalized using *CLATHRIN* as an internal standard. The specific primers used are summarized in table 1. In order to examine whether gene expression was significantly different between treatments or genotypes, ANOVA followed by a Turkey test or Student T Test's were applied.

## GUS staining

Samples were prefixed for 45 minutes at room temperature in a solution containing 50 mM NaPO4 pH7, 1,5% formaldehyde and 0,05% Triton X100. Then plants are washed 3 times in 50 mM NaPO4 pH7 with 0.05% Triton X100. Thereafter, samples were stained in 50 mM NaPO4 pH7, 0,5 mM Ferricyanide, 0,5 mM Ferrocyanide, 0,05% Triton X100 and 1 mM X-Gluc at 37°C for 48h in the dark. Afetr GUS staining, plants were washed in 50 mM NaPO4 pH7 for 5 minutes and chlorophyll was removed by EtOH increasing concentrations solutions.
The samples were mounted in 20% EtOH and were imaging by using x10 N.A. 0,3 or x40 N.A. 0,75 objectives on an Olympus BX61 microscope equipped with a camera Jenoptik Progres.

## Confocal microscopy

For confocal microscopy observation, plant samples were mounted in osmosed water and viewed on a Leica SP8 Confocal Microscope (Leica). GFP/YFP were excited at 488nm by an Argon source and re-emitted light was filtered by a passing band of 505-530nm. Laser intensity was adjusted for each line and pinhole was settled at 1 unit, resolution at 1024 x 1024 to compare expression and accumulation of reporter proteins. Quantification of fluorescence in roots was performed using Image J software. The data represent the mean pixel values in a ROI.

## RESULTS

### Regulation of auxin carriers mRNA levels in response to NO₃⁻.

To determine how auxin transporter genes respond to NO₃⁻ and whether this response is dependent on NRT1.1, we measured by quantitative RT-PCR the transcript levels of ten auxin transporters in roots of both WT and *chl1-5* (*NRT1.1* null mutant) plants grown for 8 days on either N-free medium or on medium containing 1mM KNO₃. The ten auxin transporter genes reported to contribute to auxin traffic in LRs (*PIN1-4*, *PIN7*, *ABCB1*, *ABCB4*, *ABCB19*, *AUX1* and *LAX3*) were sorted in three different groups according to their expression patterns (Figure 1). The first one is constituted by *PIN2*, *PIN3* and *ABCB1*, for which mRNA levels are not significantly influenced either by NO₃⁻ supply or by *chl1-5* mutation (Figure 1A). The second group gathers *ABCB4*, *ABCB19*, *PIN1*, *PIN4*, and *PIN7*, for which mRNA levels are modulated by NO₃⁻. Among these genes, *ABCB4* is the only one to be repressed upon NO₃⁻ supply, whereas the others are induced (Figure 1B). The NO₃⁻-responses of these genes are the same in both WT and *chl1-5* plants, suggesting that NRT1.1 plays no role in these responses. The last group contains *AUX1* and *LAX3* (Figure 1C). Both genes show higher expression in presence of NO₃⁻ as compared to N-free medium in WT plants, but the difference is suppressed or strongly attenuated in *chl1-5* mutant. This is due to the up-regulation of these two genes in the *chl1-5* mutant as compared to the WT in N-free
condition, but not on 1 mM NO3-. Therefore, NRT1.1 appears to act as a negative regulator of the expression of both AUX1 and LAX3 in the absence, but not in the presence of NO3-. Altogether, these data show that although the majority of the auxin carrier genes investigated (7 out of 10) respond to NO3- provision in the roots, only two of them display a NO3- response that seems to involve NRT1.1 as a regulator.

Protein level and localization of auxin carriers in response to NO3-.

The response to NO3- of NRT1.1 and of the ten auxin transporters was evaluated at the protein level in LRPs using reporter-tagged transgenic plants (translational fusions with GFP or YFP) in WT or chl1 mutant genetic background. Confocal sections through roots of transgenic plants grown in similar conditions to those used for the quantitative RT-PCR measurements were performed to compare protein accumulation and localization in LRPs in different conditions and at different developmental stages.

As reported previously (Bouguyon *et al.*, 2016), the protein expression of NRT1.1 in LRPs is strongly regulated by NO3-. Indeed, the GFP signal in roots of plants expressing a *NRT1.1::GFP* transgene under the control of the *NRT1.1* promoter in *chl1-5* background indicates that NRT1.1 is present in pre-emerged LRPs in the absence, but not in the presence of 1 mM NO3- in the medium (Supplemental Fig. 1). In NO3-fed plants, NRT1.1 expression is only observed in LRPs after emergence. Most auxin transporters (8 out of 10) do not show any significant NO3-dependent change either in protein accumulation or localization in the context of LRP development and emergence. In our conditions and regardless the presence of NO3-, ABCB4 and PIN4 are not detectable in LRPs and PIN7 appears to be only transiently expressed at stages I and II but then rapidly disappears as LRPs progress to next phase (data not shown). Moreover, as previously shown (Benkova *et al.*, 2003), PIN2 is not expressed in LRPs before emergence (Supplemental Fig. 2). Neither the localization nor the protein abundance of PIN1, ABCB19 and AUX1 are significantly affected by NO3- or chl1 mutation (Supplemental Figs 3, 4 and 5). Finally, although the GFP signal associated with the ABCB1-GFP fusion protein seemed slightly increased in the presence of NO3-, this was not found to be statistically significant after quantification (Supplemental Fig. 6).

In contrast, PIN3, and LAX3 are responsive to NO3- treatments. In WT plants and in presence of NO3-, PIN3-GFP spatio-temporal expression pattern in LRPs is similar to that previously
shown (Benkova *et al.*, 2003). Indeed, PIN3 is expressed from the first developmental stage to emergence and its early widespread localization is progressively restricted to the meristematic zone (Figure 2A- a-c right panels). However, on N-free medium, PIN3-GFP signal becomes undetectable in LRPs from stage IV-V up to emergence (Figure 2A b-c left panel). To determine whether this is due to a NO₃⁻ specific effect, PIN3-GFP expression was investigated in plants grown on 0.5 mM glutamine as an alternative N source (Figure 2A, central panels). Supply of glutamine results in a PIN3-GFP expression similar to that recorded with NO₃⁻, indicating that lack of expression on N-free medium is not due to the absence of NO₃⁻ *per se*, but most probably results from N-deficiency. In agreement with the hypothesis that PIN3-GFP is not specifically regulated by NO₃⁻, the effect of N on this protein is NRT1.1-independent since the same pattern is observed in a WT or a *chl1* mutant background (Figure 2B, 2C).

As previously shown (Swarup *et al.*, 2008), LAX3-YFP signal in WT background is detected from stage II to stage VII in mature stele and in cortical or epidermal cells overlaying LRPs under NO₃⁻ supply (Figure 3A- a to c right panels). However, LAX3-YFP signal is strongly decreased in cells overlying LRPs, regardless of the LRP developmental stage, in response to either N-starvation (Figure 3A- a to c left panel) or supply of glutamine as a N source (Figure 3A, middle panel). Interestingly, this down-regulation of LAX3-YFP expression is not recorded in *chl1-5* mutant background (Figure 3B, 3C). These data show that NRT1.1 acts as a negative regulator of LAX3 expression in the absence of NO₃⁻. Interestingly, the *chl1-5* mutation restores LAX3-YFP signal in LRP overlying cells only after LRPs have reached stage IV (Figure 3A and B-compare a with b and c stages), indicating that NRT1.1 is responsible for LAX3 extinction at late, but not at early stages of LRP development. This is fully consistent with the observation that NRT1.1 starts repressing LRP development at stages IV/V (Krouk *et al.*, 2010; Bouguyon *et al.*, 2016). In conclusion, the pattern of LAX3-YFP expression around LRPs in WT and *chl1* mutant background is in agreement with that of *LAX3* transcript accumulation in the roots (i.e., both are up-regulated by NO₃⁻ in WT plants and by *chl1* mutation in the absence of NO₃⁻).

It has been reported that the upregulation of *LAX3* expression that facilitates LRP outgrowth is triggered by the LBD29 transcription factor (Swarup *et al.*, 2008; Porco *et al.*, 2016). To test whether LBD29 responds to NO₃⁻ in a NRT1.1-dependent manner, its transcript accumulation in roots was investigated in roots of Col or *chl1-5* plants grown on N-free
medium or on 1 mM NO3-. The data indicate that expression of *LBD29* is strongly induced by NO3- supply as compared to N-free conditions in WT plants, and that it is markedly overexpressed in the *chl1-5* mutant as compared with the WT on N-free medium, but not in the presence of NO3- (Figure 3D). This is fully consistent with the expression patterns of both *LAX3* gene (Figure 1) and LAX3-GFP protein (Figure 3), which therefore suggests that LAX3 response to NO3- is mediated by the NRT1.1-dependent regulation of *LBD29* expression.

**The TAR2 auxin biosynthesis enzyme is up-regulated by NO3- in a NRT1.1-dependent manner.**

The *TAR2* gene was reported to be N-responsive and to contribute to the regulation of LR development by NH4NO3 supply (Ma *et al.*, 2014). However, its putative role in the specific response to NO3- in connection with the NRT1.1-dependent pathway is not known. Therefore, we first investigated the effect of the three N regimes (N-free, 0.5 mM glutamine or 1 mM NO3-) on *TAR2* mRNA accumulation in the roots of WT and *chl1-5* plants, as well as in those of the *tar2c* knock-out mutant of *TAR2*. Although *TAR2* mRNA accumulation was previously shown to be down-regulated by increasing NH4NO3 concentrations (Ma *et al.*, 2014), it was in our conditions markedly induced in the WT by N supply as compared with N-free condition, with a much more pronounced response on NO3- as compared with glutamine (Figure 4A). This NO3- induction was fully lost in the *chl1-5* mutant because *TAR2* expression was already high in this line on N-free medium or on gln. As expected, no significant *TAR2* mRNA level was recorded in the *tar2c* knock-out mutant.

Both the induction of *TAR2* expression by NO3-, and its up-regulation in N-starved *chl1-5* plants as compared to WT were confirmed by histological staining of GUS activity in *pTAR2::GUS* plants (Figure 4B). As shown previously (Ma *et al.*, 2014), *TAR2* expression in roots is predominantly localized in the stele, and we did not observe *pTAR2* activity in endodermis, cortex or epidermis (Figure 4B and 4C). In presence of either 0.5 mM gln or 1 mM NO3-, strong GUS staining was also observed inside the vast majority (>90%) of young LRPs (stages I-III) in both WT and *chl1-5* plants (Figure 4C). On N-free medium, GUS staining in LRPs was only visible in ca~60% of the young LRPs of *chl1-5* plants, and at a lower intensity than that recorded in plants fed with either gln or NO3- (Figure 4C). In all cases, *pTAR2::GUS* expression in LRPs was no more recorded as soon as these LRPs
evolved to later stages of development (stages IV-V onwards). Altogether, the above results show that in the absence of NO3-, NRT1.1 acts as a strong negative regulator of TAR2 expression in the stele and to a lower extent in the young LRPs. This indicates that NRT1.1 may control LRP development not only through its auxin transport facilitation within the LRPs (Krouk et al., 2010), but also by modulating auxin biosynthesis in the stele. In addition, these results also show that TAR2 expression may not only be responsive to NRT1.1-mediated NO3- signaling but also to another N stimulatory effect that can be activated by gln supply.

TAR2 is required for the stimulation of LR development by NO3- and its regulation by
NRT1.1

The induction of *TAR2* expression in the roots by NO3⁻ suggests that increased auxin biosynthesis is associated with NO3⁻ regulation of RSA. To determine whether TAR2 may contribute to the NO3⁻ stimulation of LR development by locally increasing auxin provision to LRPs, expression of *DR5::GFP* was investigated in both WT and *tar2c* mutant backgrounds in the roots of plants grown on 1 mM NO3⁻ (Figure 5). The GFP signal was predominantly found in the stele and LRPs of both genotypes, but with a strongly reduced intensity in *tar2c* as compared to WT (Figure 5). The negative effect of *TAR2* mutation on *DR5::GFP* expression in LRPs was found at almost all stages of development, with the exception of stage VII, just prior to emergence (Figures 5B and 5C). This indicates that TAR2-dependent auxin biosynthesis in the stele and/or the LRPs is required for increasing auxin accumulation/signaling in LRPs in response to NO3⁻. Note however that although important, the presence of a functional *TAR2* gene is not strictly required to generate *DR5::GFP* expression in the root. This expression is still significant in the stele and in LRPs at all developmental stages in the *tar2c* mutant (Figures 5A, 5B and 5C), suggesting that auxin partly comes from another origin than the TAR2-dependent biosynthesis pathway.

TAR2 mutation alone (*tar2c* mutant) had only very limited effect, if any, on the growth of the primary root in the three conditions tested (Figures 6A and 6B). However, it very significantly decreased (by nearly 50%) the density of LRs in the plants grown on NO3-, but not in those supplied with either glutamine of N-free medium (Figures 6A and 6C). This demonstrates that TAR2 participates in the stimulation of LR development by NO3-, in agreement with the observations that TAR2 expression is induced by NO3- and that its
mutation lowers *DR5* activity in LRPs (Figures 4 and 5). Combining *NRT1.1* and *TAR2* mutations (double *chl1-5tar2c* mutant) had no additional effect on LR development as compared to the single *TAR2* mutation, but strikingly affected it as compared to the single *NRT1.1* mutation (Figures 6A and 6C). Indeed, the dramatic increase of LR density classically observed in *chl1-5* as compared to the WT on NO₃⁻-free medium (Krouk *et al.*, 2010; Bouguyon *et al.*, 2015) was totally abolished in the double *chl1-5tar2c* mutant. These data indicate that the up-regulation of *TAR2* expression in response to *NRT1.1* mutation (Figure 4) is most likely one key response required to trigger enhanced LR development in *chl1-5* plants.

DISCUSSION

Nitrate regulation of auxin carriers in lateral root primordia

One striking conclusion arising from our work is that, with the noticeable exception of LAX3, NRT1.1 has little effect on the expression of the auxin carriers involved in LRP development. However, as suggested by previous reports on *Arabidopsis* (Li *et al.*, 2011) and rice (Naz *et al.*, 2019; Sun *et al.*, 2017), our data show that the root expression of key auxin carriers is responsive to N supply in WT plants. Although not dramatic, changes in mRNA levels in the whole root were found in response to NO₃⁻ for 7 out of the 10 auxin carriers investigated (Figure 1). When considering more precisely the expression at the protein level in or around the LRPs of WT background plants, only 2 of them (namely, PIN3 and LAX3) displayed a visible increase in the fluorescence from GFP or YFP fusion proteins on 1 mM NO₃⁻ as compared to N-free medium (Figures 2 and 3). Most importantly, for PIN3 this effect was not specific of NO₃⁻ since the same increase in fluorescence was observed upon provision of 0.5 mM gln as a N source (Figure 2). This suggests that the response to NO₃⁻ of PIN3 is not specifically due to NO₃⁻ signaling, but either to a signaling action of an N metabolite downstream of NO₃⁻ assimilation, or to more general effect of N nutrition. As a matter of fact, organic forms of N, and in particular amino acids, have been shown to specifically control several mechanisms involved in root growth and development in *Arabidopsis* (Walch-Liu *et al.*, 2006; Gifford *et al.*, 2008; Vidal *et al.*, 2010). Our data indicate that none of the auxin carriers expressed in LRPs (PIN1-4, PIN7, ABCB1, ABCB4, ABCB19 and AUX1) displays a regulation pattern correlated with that previously reported for *DR5* activity in LRPs (e.g., repression in the absence of NO₃⁻ that is suppressed in the
chl1-5 mutant, (see Krouk et al., 2010). This supports the hypothesis that NRT1.1 modulates auxin accumulation in LRP directly through its auxin transport capacity, and not indirectly through modified expression of other auxin carriers. However, it cannot be ruled out that NRT1.1 may act on these carriers at a regulatory level that was not investigated in our study, by triggering posttranslational modifications affecting transport activity for instance. Furthermore, although not predominantly expressed in LRPs, ABCB21 has recently been reported to markedly affect DR5 activity in these organs (Jenness et al., 2019), but its regulation by NO3- is unknown. Therefore, further investigation is needed to conclude whether the above listed auxin carriers are regulated by NRT1.1, or not.. In contrast, our data on LAX3 mRNA accumulation and of LAX3-YFP fusion protein expression clearly indicate that the expression of this carrier in overlying tissues of the LRPs is under control by NRT1.1-dependent NO3- signaling. First, the stimulation by NO3- of LAX3-YFP expression in cortical cells cannot be mimicked by gln supply (Figure 3). Second, both the increases in LAX3 gene and LAX3-YFP protein expression in response to NO3- were suppressed in the chl1-5 mutant (Figures 1 and 3). This is further supported by the fact that the gene expression of LBD29, the direct positive regulator of LAX3 (Porco et al., 2016), displays the same pattern of regulation (Figure 3). In agreement with what was previously recorded for LR density and for auxin accumulation/signaling in LRP (DR5 activity), the stimulatory effect of NO3- was suppressed because LAX3 and LBD29 expression were already high in the chl1-5 mutant in the absence of NO3-, indicating a negative regulatory role of NRT1.1 under this condition (Figures 1 and 3). The mechanisms by which NRT1.1 may lead to repression of LBD29 and LAX3 expression are not known. However, the stimulation of both LBD29 and LAX3 expression in cells overlying LRPs was proposed to be due to the induction of LBD29 and LAX3 genes by the auxin diffusing from the LRP to the overlying tissues (Swarup et al., 2008; Porco et al., 2016). Therefore, it is conceivable that the NRT1.1-mediated repression of auxin accumulation in LRPs in the absence of NO3- (Krouk et al., 2010; Bouguyon et al., 2015; Bouguyon et al., 2016; Zhang et al., 2019), may lead to a lowered auxin diffusion out of the LRPs, which may in turn limit induction of both LBD29 and LAX3 genes.
Auxin biosynthesis is a target of NRT1.1-dependent NO₃⁻ signaling in roots

The importance of local auxin biosynthesis for controlling plant development has been
recently reconsidered, and shown to be important for responses to internal or external signals
(ethylene, light, temperature and pathogens, Ljung et al., 2001; Brumos et al., 2018; Zhao et
al., 2018). Data concerning RSA responses to nutrients are scarce on this point (Guan et al.,
2017; Olatunji et al., 2017), with the exception of the report by Ma et al., (2014), indicating
that the TAR2 auxin biosynthetic gene contributes to the root foraging response triggered by
mild N-limitation in Arabidopsis. This response is considered to be due to a repression
exerted by high N supply, which is not specific to NO3-, and may involve systemic signaling
(Forde, 2014; Giehl and von Wirén, 2014). Our data confirm a key role of TAR2 in the N
regulation of LR growth, but in a different context that is the local stimulation of LRP
development by NO3- signaling. Indeed, TAR2 expression is induced by NO3- (Figure 4), and
tar2 mutants display a markedly reduced increase in LR density in response to NO3- supply
as compared with WT plants (Figure 6). This is correlated with reduced DR5 activity in both
the stele and LRPs of plants with tar2 as compared to WT background (Figure 5). This
suggests that the TAR2 biosynthetic pathway is an important source of auxin for ensuring
proper accumulation of the hormone in LRPs and for yielding full stimulation of LR
development by NO3-. However, because neither DR5 activity nor LR emergence is totally
suppressed in the tar2 mutant, it is likely that another auxin source, coming from acropetal
transport from the shoot, other biosynthesis and/or conjugation pathway (Bhalerao et al.,
2002; Petrasek and Friml, 2009; Olatunji et al., 2017), also contributes to supply auxin to the
LRPs. As it is the case for LR density and key factors associated with it (DR5 activity in
LRPs, LBD29/LAX 3 expression), the stimulation of TAR2 expression by NO3- in the stele is
actually mostly due to the relief of the repression exerted by NRT1.1 in the absence of NO3-.
This is illustrated by the upregulation of TAR2 in the chl1-5 mutant on N-free medium
(Figure 4). This TAR2 upregulation appears to be crucial for the strongly increased LR
growth phenotype of the chl1-5 plants because the double chl1-5tar2c mutant did not differ in
LR density from the WT in NO3- free media (Figure 4). The mechanism associated with the
NRT1.1-dependent repression of TAR2 expression in the stele in the absence of NO3- is
unclear. One hypothesis is that the stimulation of basipetal auxin transport out of the LRPs by
NRT1.1 (Krouk et al., 2010) may locally increase auxin levels in the stele. Because the
TAR2 auxin biosynthetic pathway has been recently reported to be feedback repressed by
auxin (Suzuki et al., 2015; Takato et al., 2017), this may explain why TAR2 expression is
reduced in the stele of WT plants deprived of NO3⁻ (Figure 4). Alternatively, because NRT1.1
has been shown to be also expressed in the central cylinder of mature portions of the PR,
where it contributes to xylem loading of NO3⁻ (Léran *et al.*, 2013), it is also possible that
NRT1.1 may act on TAR2 expression directly in the stele when NO3⁻ is absent from the
medium.

Interestingly, *TAR2* was found to be expressed in a N-stimulated manner in young LRPs at stages I-III (Figure 4C), *i.e.*, at earlier developmental stages than those (IV-V) where NRT1.1 starts to act as a regulator of LRP emergence (Bouguyon *et al.*, 2016). Because *TAR2* expression in young LRPs is responsive to both gln and NO3⁻, this suggests that N can modulate early LRP development by controlling auxin biosynthesis *in situ* independently of NRT1.1. This is in line with the previous proposal that N can act on various developmental check points of LR development through different mechanisms (Forde, 2014; Giehl and von Wirén, 2014).

A more integrated model for NRT1.1-dependent regulation of LR development by NO3⁻.

Our previous working model for explaining the NRT1.1-dependent repression of LR growth
in the absence or at low availability (<0.5 mM) of NO3⁻ almost entirely relied on the
hypothesis that NRT1.1 inhibits LRP outgrowth (more specifically at developmental stage
IV) because it facilitates the basipetal auxin transport out of the LRP, therefore preventing the
hormone accumulation at the LRP tip (Krouk *et al.*, 2010; Bouguyon *et al.*, 2015). This
hypothesis was supported by the auxin transport capacity of NRT1.1 recorded in
heterologous systems (Krouk *et al.*, 2010; Bouguyon *et al.*, 2015; Wei *et al.*, 2018; Zhang *et
al.*, 2019), and by its largely predominant localization in the outer cell layer of the LRP
(Krouk *et al.*, 2010), which corresponds to the basipetal auxin transport route (Benkova *et al.*,
2003). Because increasing NO3⁻ concentrations (>0.5 mM) results both in an inhibition of the
NRT1.1-mediated auxin transport facilitation (Krouk *et al.*, 2010) and in a repression of the
NRT1.1 protein expression in LRPs (Bouguyon *et al.*, 2016), we proposed that the local
stimulation of LR growth by NO3⁻ is due to a relief of the repression exerted by NRT1.1. The
data reported here strengthen our previous conclusions, but show that NRT1.1 affects other
processes in addition to auxin distribution within the LRP, such as the biosynthesis of the
hormone and the mechanical resistance of the PR tissues to LRP outgrowth. As summarized
in Figure 7, the repressive role of NRT1.1 on LRP development can now be extended to three
separate but synergistic actions. Indeed, NRT1.1 prevents auxin accumulation in the LRP not
only by favoring the basipetal transport of the hormone, but also by triggering the repression
of *TAR2* expression in the stele. As a consequence of reduced auxin biosynthesis, it is likely
that the acropetal flux channeling the hormone to the LRP tip is also lowered, therefore
contributing to slowing down LRP development and emergence. In addition, NRT1.1 acts as
a negative regulator of the expression of LAX3 in the PR tissues overlying the LRPs. Given
the proposed function of LAX3 in LRP development (Swarup *et al.*, 2008), this is expected to
prevent the cell wall relaxation in the LRP overlying tissues that is required to lower the
mechanical resistance to LRP emergence (Lucas *et al.*, 2013).

Beyond the repression of LRP development at low NO3- availability summarized in Figure 7, NRT1.1 has also been reported to favor LR elongation in NO3- -rich patches of the external medium (Remans et al., 2006), and to trigger the induction of the *AFB3* auxin receptor gene by NO3- that results in repressed PR growth and stimulated LR initiation (Vidal et al., 2010; Vidal et al., 2014). By interfering with auxin biosynthesis, transport, signaling, and thus by controlling various key checkpoints of LR development, our studies reveal that NRT1.1 has a much more integrated function in governing the NO3- regulation of RSA in *Arabidopsis* than previously anticipated (Vidal et al., 2014; Bouguyon et al., 2015). It may therefore be expected that all aspects and mechanisms of this integrated function are not fully unraveled yet.
ACKNOWLEDGMENTS

P.N., G.K., E.Ba., L.B. and A.G. designed and supervised the experiments; A.M., E.Bo., C.C.M., F.P.W., P.N. and L.B. performed experiments; A.M. and E.Bo. generated plant material; A.M., E.BO., C.C.M., G.K., P.N., E.Ba., L.B. and A.G. analyzed the data , A.M., L.B., and A.G. wrote the article with contribution of all the authors.

This work was supported by the Agence Nationale de la Recherche program "Programme PCRI ANR-16-CE20-0022". We would like to thank members of the lab for discussion. We are grateful to Yiping Tong for the gift of tar2-c (SALK_143208.56.00.x) and TAR2pro::GUS (CS16434) seeds from the Arabidopsis Bio-logical Resource Center (ABRC). We thank the Montpellier Rio Imaging platform for microscopy observations.
BIBLIOGRAPHY

Bailly A, Sovero V, Vincenzetti V, Santelia D, Bartnik D, Koenig BW, Mancuso S, Martinoia E, Geisler M. 2008. Modulation of P-glycoproteins by auxin transport inhibitors is mediated by interaction with immunophilins. Journal of Biological Chemistry **283**, 21817-21826.

Bellegarde F, Gojon A, Martin A. 2017. Signals and players in the transcriptional regulation of root responses by local and systemic N signaling in Arabidopsis thaliana. Journal of Experimental Botany **68**, 2553-2565.

Benkova E, Michniewicz M, Sauer M, Teichmann T, Seifertova D, Jurgens G, Friml J. 2003. Local, efflux-dependent auxin gradients as a common module for plant organ formation. Cell **115**, 591-602.

Bhalerao RP, Eklof J, Ljung K, Marchant A, Bennett M, Sandberg G. 2002. Shoot-derived auxin is essential for early lateral root emergence in Arabidopsis seedlings. The Plant Journal **29**, 325-332.

Blilou I, Xu J, Wildwater M, Willemsen V, Paponov I, Friml J, Heidstra R, Aida M, Palme K, Scheres B. 2005. The PIN auxin efflux facilitator network controls growth and patterning in Arabidopsis roots. Nature **433**, 39-44.

Bouguyon E, Brun F, Meynard D, et al 2015. Multiple mechanisms of nitrate sensing by Arabidopsis nitrate transceptor NRT1.1. Nature Plants **1**, 15015.

Bouguyon E, Perrine-Walker F, Pervent M, et al. 2016. Nitrate Controls Root Development through Posttranscriptional Regulation of the NRT1.1/NPF6.3 Transporter/Sensor. Plant Physiology **172**, 1237-1248.

Brumos J, Robles LM, Yun J, Vu TC, Jackson S, Alonso JM, Stepanova AN. 2018. Local Auxin Biosynthesis Is a Key Regulator of Plant Development. Developmental Cell **47**, 306-318.e305.

Brunoud G, Wells DM, Oliva M, et al. 2012. A novel sensor to map auxin response and distribution at high spatio-temporal resolution. Nature **482**, 103-106.

Cho M, Lee SH, Cho HT. 2007. P-Glycoprotein4 Displays Auxin Efflux Transporter-Like Action in Arabidopsis Root Hair Cells and Tobacco Cells. The Plant Cell Online **19**, 3930-3943.

Crawford NM, Glass ADM. 1998. Molecular and physiological aspects of nitrate uptake in plants. Trends in Plant Science **3**, 389-395.

Drew MC. 1975. Comparison of the effects of a localised supply of phosphate, nitrate, ammonium and potassium on the growth of the seminal root system, and the shoot in barley. New Phytologist **75**, 479-490.

Dubrovsky JG, Sauer M, Napsucialy-Mendivil S, Ivanchenko MG, Friml J, Shishkova S, Celenza J, Benkova E. 2008. Auxin acts as a local morphogenetic trigger to specify lateral root founder cells. Proc Natl Acad Sci U S A **105**, 8790-8794.

Forde BG. 2002. Local and long-range signaling pathways regulating plant responses to nitrate. Annual Reviews Plant Biology **53**, 203-224.

Forde BG. 2014. Nitrogen signalling pathways shaping root system architecture: an update. Current Opinion in Plant Biology **21**, 30-36.

Fukaki H, Okushima Y, Tasaka M. 2007. Auxin-mediated lateral root formation in higher plants. International Review Cytology **256**, 111-137.

Giehl RF, von Wiren N. 2014. Root nutrient foraging. Plant Physiology **166**, 509-517.
Giehl RFH, Lima JE, von Wirén N. 2012. Localized iron supply triggers lateral root elongation in Arabidopsis by altering the AUX1-mediated auxin distribution. The Plant Cell **24**, 33-49.

Gifford ML, Dean A, Gutierrez RA, Coruzzi GM, Birnbaum KD. 2008. Cell-specific nitrogen responses mediate developmental plasticity. Proc Natl Acad Sci U S A **105**, 803-808.

Gojon A, Krouk G, Perrine-Walker F, Laugier E. 2011. Nitrate transceptor(s) in plants. Journal of Experimental Botany **62**, 2299-2308.

Gojon A, Nacry P, Davidian JC. 2009. Root uptake regulation: a central process for NPS homeostasis in plants. Current Opinion in Plant Biology **12**, 328-338.

Guan P. 2017. Dancing with Hormones: A Current Perspective of Nitrate Signaling and Regulation in Arabidopsis. Frontiers in Plant Science **8**, 1697.

Gutierrez RA, Lejay LV, Dean A, Chiaromonte F, Shasha DE, Coruzzi GM. 2007. Qualitative network models and genome-wide expression data define carbon/nitrogen-responsive molecular machines in Arabidopsis. Genome Biology **8**, R7.

Hermans C, Hammond JP, White PJ, Verbruggen N. 2006. How do plants respond to nutrient shortage by biomass allocation? Trends in Plant Science **11**, 610-617.

Ho CH, Lin SH, Hu HC, Tsay YF. 2009. CHL1 functions as a nitrate sensor in plants. Cell **138**, 1184-1194.

Jacquot A, Li Z, Gojon A, Schulze W, Lejay L. 2017. Post-translational regulation of nitrogen transporters in plants and microorganisms. Journal of Experimental Botany **68**, 2567-2580.

Jenness MK, Carraro N, Pritchard CA, Murphy AS. 2019. The Arabidopsis ATP-BINDING CASSETTE Transporter ABCB21 regulates auxin levels in cotyledons, the root pericycle, and leaves. Frontiers in Plant Science **10**, 806.

Kiba T, Krapp A. 2016. Plant Nitrogen Acquisition Under Low Availability: Regulation of Uptake and Root Architecture. Plant and Cell Physiology **57**, 707-714.

Krouk G, Lacombe B, Bielach A, et al. 2010. Nitrate-regulated auxin transport by NRT1.1 defines a mechanism for nutrient sensing in plants. Developmental Cell **18**, 927-937.

Kubes M, Yang H, Richter GL, et al. 2012. The Arabidopsis concentration-dependent influx/efflux transporter ABCB4 regulates cellular auxin levels in the root epidermis. The Plant Journal **69**, 640-654.

Leran S, Munos S, Brachet C, Tillard P, Gojon A, Lacombe B. 2013. Arabidopsis NRT1.1 is a bidirectional transporter involved in root-to-shoot nitrate translocation. Molecular Plant **6**, 1984-1987.

Li B, Li Q, Su Y, Chen H, Xiong L, Mi G, Kronzucker HJ, Shi W. 2011. Shoot-supplied ammonium targets the root auxin influx carrier AUX1 and inhibits lateral root emergence in Arabidopsis. Plant, Cell and Environment **34**, 933-946.

Liu KH, Tsay YF. 2003. Switching between the two action modes of the dual-affinity nitrate transporter CHL1 by phosphorylation. The EMBO Journal **22**, 1005-1013.

Ljung K, Bhalerao RP, Sandberg G. 2001. Sites and homeostatic control of auxin biosynthesis in Arabidopsis during vegetative growth. The Plant Journal **28**, 465-474.

Lucas M, Kenobi K, von Wangenheim D, et al. 2013. Lateral root morphogenesis is dependent on the mechanical properties of the overlaying tissues. Proc Natl Acad Sci U S A **110**, 5229-5234.

Ma W, Li J, Qu B, He X, Zhao X, Li B, Fu X, Tong Y. 2014. Auxin biosynthetic gene TAR2 is involved in low nitrogen-mediated reprogramming of root architecture in Arabidopsis. The Plant Journal **78**, 70-79.

Malamy J, Benfey P. 1997. Organization and cell differentiation in lateral roots of Arabidopsis thaliana. Development (Cambridge, England) **124**, 33-44.
Marchant A, Bhalerao R, Casimiro I, Eklöf J, Casero PJ, Bennett M, Sandberg G. 2002. AUX1 promotes lateral root formation by facilitating indole-3-acetic acid distribution between sink and source tissues in the Arabidopsis seedling. The Plant Cell **14**, 589-597.

Miller AJ, Fan X, Orsel M, Smith SJ, Wells DM. 2007. Nitrate transport and signalling. Journal of Experimental Botany **58**, 2297-2306.

Mounier E, Pervent M, Ljung K, Gojon A, Nacry P. 2014. Auxin-mediated nitrate signalling by NRT1.1 participates in the adaptive response of Arabidopsis root architecture to the spatial heterogeneity of nitrate availability. Plant, Cell and Environment **37**, 162-174.

Mravec J, Kubes M, Bielach A, Gaykova V, Petrasek J, Skupa P, Chand S, Benkova E, Zazimalova E, Friml J. 2008. Interaction of PIN and PGP transport mechanisms in auxin distribution-dependent development. Development **135**, 3345-3354.

Munos S, Cazettes C, Fizames C, Gaymard F, Tillard P, Lepetit M, Lejay L, Gojon A. 2004. Transcript profiling in the chl1-5 mutant of Arabidopsis reveals a role of the nitrate transporter NRT1.1 in the regulation of another nitrate transporter, NRT2.1. The Plant Cell **16**, 2433-2447.

Nacry P, Bouguyon E, Gojon A. 2013. Nitrogen acquisition by roots: physiological and developmental mechanisms ensuring plant adaptation to a fluctuating resource. Plant and Soil **370**, 1-29.

Naz M, Luo B, Guo X, Li B, Chen J, Fan X. 2019. Overexpression of Nitrate Transporter OsNRT2.1 Enhances Nitrate-Dependent Root Elongation. Genes (Basel) **10**.

O'Brien JA, Vega A, Bouguyon E, Krouk G, Gojon A, Coruzzi G, Gutierrez RA. 2016. Nitrate Transport, Sensing, and Responses in Plants. Molecular Plant **9**, 837-856.

Olatunji D, Geelen D, Verstraeten I. 2017. Control of Endogenous Auxin Levels in Plant Root Development. International Journal of Molecular Sciences **18**.

Petrasek J, Friml J. 2009. Auxin transport routes in plant development. Development **136**, 2675-2688.

Porco S, Larrieu A, Du Y, et al. 2016. Lateral root emergence in Arabidopsis is dependent on transcription factor LBD29 regulation of auxin influx carrier LAX3. Development **143**, 3340-3349.

Remans T, Nacry P, Pervent M, Filleur S, Diatloff E, Mounier E, Tillard P, Forde BG, Gojon A. 2006. The Arabidopsis NRT1.1 transporter participates in the signaling pathway triggering root colonization of nitrate-rich patches. Proc Natl Acad Sci U S A **103**, 19206-19211.

Ruffel S, Krouk G, Ristova D, Shasha D, Birnbaum KD, Coruzzi GM. 2011. Nitrogen economics of root foraging: transitive closure of the nitrate-cytokinin relay and distinct systemic signaling for N supply vs. demand. Proc Natl Acad Sci U S A **108**, 18524-18529.

Santelia D, Vincenzetti V, Azzarello E, Bovet L, Fukao Y, Duchtig P, Mancuso S, Martinoia E, Geisler M. 2005. MDR-like ABC transporter AtPGP4 is involved in auxin-mediated lateral root and root hair development. FEBS Letters **579**, 5399-5406.

Sun CH, Yu JQ, Hu DG. 2017. Nitrate: A Crucial Signal during Lateral Roots Development. Frontiers in Plant Science **8**, 485.

Suzuki M, Yamazaki C, Mitsui M, Kakei Y, Mitani Y, Nakamura A, Ishii T, Soeno K, Shimada Y. 2015. Transcriptional feedback regulation of YUCCA genes in response to auxin levels in Arabidopsis. The Plant Cell Reports **34**, 1343-1352.

Swarup K, Benková E, Swarup R, et al. 2008. The auxin influx carrier LAX3 promotes lateral root emergence. Nature Cell Biology **10**, 946-954.

Swarup R, Kargul J, Marchant A, et al. 2004. Structure-function analysis of the presumptive Arabidopsis auxin permease AUX1. The Plant Cell **16**, 3069-3083.

Swarup R, Peret B. 2012. AUX/LAX family of auxin influx carriers-an overview. Frontiers in Plant Science **3**, 225.
Takato S, Kakei Y, Mitsui M, et al. 2017. Auxin signaling through SCF(TIR1/AFBs) mediates feedback regulation of IAA biosynthesis. Bioscience, biotechnology, and biochemistry **81**, 1-7.

Tsay YF, Schroeder JI, Feldmann KA, Crawford NM. 1993. The herbicide sensitivity gene CHL1 of Arabidopsis encodes a nitrate-inducible nitrate transporter. Cell **72**, 705-713.

Ulmasov T, Murfett J, Hagen G, Guilfole TJ. 1997. Aux/IAA proteins repress expression of reporter genes containing natural and highly active synthetic auxin response element. The Plant Cell **9**, 1963-1971

Vidal EA, Alvarez JM, Gutierrez RA. 2014. Nitrate regulation of AFB3 and NAC4 gene expression in Arabidopsis roots depends on NRT1.1 nitrate transport function. Plant Signaling and Behavior **9**, e28501.

Vidal EA, Araus V, Lu C, Parry G, Green PJ, Coruzzi GM, Gutierrez RA. 2010. Nitrate-responsive miR393/AFB3 regulatory module controls root system architecture in Arabidopsis thaliana. Proc Natl Acad Sci U S A **107**, 4477-4482.

Vieten A, Sauer M, Brewer PB, Friml J. 2007. Molecular and cellular aspects of auxin-transport-mediated development. Trends in Plant Science **12**, 160-168.

Vieten A, Vanneste S, Wisniewska J, Benkova E, Benjamins R, Beeckman T, Luschnig C, Friml J. 2005. Functional redundancy of PIN proteins is accompanied by auxin-dependent cross-regulation of PIN expression. Development **132**, 4521-4531.

Walch-Liu P, Forde BG. 2008. Nitrate signalling mediated by the NRT1.1 nitrate transporter antagonises L-glutamate-induced changes in root architecture. The Plant Journal **54**, 820-828.

Walch-Liu P, Liu LH, Remans T, Tester M, Forde BG. 2006. Evidence that L-glutamate can act as an exogenous signal to modulate root growth and branching in Arabidopsis thaliana. Plant and Cell Physiology **47**, 1045-1057.

Wang R, Xing X, Wang Y, Tran A, Crawford NM. 2009. A genetic screen for nitrate regulatory mutants captures the nitrate transporter gene NRT1.1. Plant Physiology **151**, 472-478.

Wei J, Zheng Y, Feng H, Qu H, Fan X, Yamaji N, Ma JF, Xu G. 2018. OsNRT2.4 encodes a dual-affinity nitrate transporter and functions in nitrate-regulated root growth and nitrate distribution in rice. Journal of Experimental Botany **69**, 1095-1107.

Xu J, Scheres B. 2005. Dissection of Arabidopsis ADP-RIBOSYLATION FACTOR 1 function in epidermal cell polarity. The Plant Cell **17**, 525-536.

Zadnikova P, Petrasek J, Marhavy P, et al. 2010. Role of PIN-mediated auxin efflux in apical hook development of Arabidopsis thaliana. Development **137**, 607-617.

Zhang H, Forde BG. 1998. An Arabidopsis MADS box gene that controls nutrient-induced changes in root architecture. Science **279**, 407-409.

Zhang X. Cui Y, Yu M, Su B, Gong W, Baluska F, Komis G, Samaj J, Lin J. 2019. Phosphorylation-Mediated Dynamics of Nitrate Transceptor NRT1.1 Regulate Auxin Flux and Nitrate Signaling in Lateral Root Growth. Plant Physiology **181**, 480-498.

Zhao Y. 2018. Essential Roles of Local Auxin Biosynthesis in Plant Development and in Adaptation to Environmental Changes. Annual Review of Plant Biology **69**, 417-435.
FIGURE LEGENDS

**Fig. 1: Relative mRNA accumulation of auxin transporters in roots of WT (Col) and chl1-5 seedlings grown in presence or absence of NO3-**

Wild-type and *chl1-5* seedlings were grown for 8 days in vertical agar plates on basal medium supplemented with 1 mM NO3- or not. Transcript levels were quantified in the roots by quantitative RT-PCR using *CLATHRIN* as an internal standard. The ten auxin transporter genes investigated are grouped into three different classes, according to the response of their mRNA levels to NO3- and *NRT1.1* mutation:

A) Auxin transporters not regulated by NO3-

B) Auxin transporters regulated by NO3- independently of *NRT1.1*.

C) Auxin transporters regulated by NO3- in a *NRT1.1*-dependent way.

The data are the mean ± se of 2 replicates from 3 independent experiments.

Statistically significant differences between genotypes or N treatments at P < 0.05 are shown by different letters and ANOVA followed by a Turkey Test were used to show these significant differences.

**Fig. 2: PIN3-GFP expression in lateral root primordia is repressed by N starvation both in WT (Col) and chl1-5 genetic backgrounds.**

Seedlings expressing PIN3-GFP in Col (A) or in *chl1-5* (B) genetic background were grown for 8 days in vertical agar plates on basal medium deprived of N (left panel) or supplemented with either 0.5mM glutamine (Gln; middle panel) or 1mM NO3- (right panel). Primary root portions with lateral root primordia at various developmental stages were observed under confocal microscope.

a = stage I-III, b = stage IV-V, c = stage VI-VII. White bar represents 100µm. The pictures shown are representative of >20 primordia from >10 plants of 3 independent experiments.

(C) Fluorescence quantification in different root tissues of 8-day-old plants expressing PIN3-GFP in *WT* or *chl1.5* lines in the absence or presence of NO3- or glutamine (Gln). Fluorescence was measured in unmerged primordia stage IV to VIII, and emerged primordia. Values are the mean of 5–10 plants from 2 independent experiments. P values were analyzed by Student's T Test and differences with N treatments are statistically significant at ***P < 0.001.
**Fig. 3: LAX3-YFP expression in tissues overlaying lateral root primordia is repressed in the absence of NO₃⁻ in a NRT1.1-dependent manner.**

Seedlings expressing LAX3-YFP in Col (A) or in *chl1-5* (B) genetic background were grown for 8 days in vertical agar plates on basal medium deprived of N (left panel) or supplemented with either 0.5mM glutamine (middle panel) or 1mM NO₃⁻ (right panel). Primary root portions with lateral root primordia at various developmental stages were observed under confocal microscope.

a = stage I-III, b = stage IV-V, c = stage VI-VII. White bar represents 100µm. The pictures shown are representative of >20 primordia from >10 plants of 3 independent experiments.

(C) Fluorescence quantification in different root tissues of 8-day-old plants expressing LAX3-YFP in *WT* or *chl1.5* lines in the absence or presence of NO₃⁻ or glutamine (Gln). Fluorescence was measured in cortical/epidermal cells surrounding unmerged primordia stage IV to VIII. Values are the mean of 5–10 plants from 2 independent experiments. P values were analyzed by Student's T Test and differences with N treatments are statistically significant at ****P < 0.0001.(D) Relative mRNA accumulation of LBD29 in roots of Col and *chl1-5* plants grown for 12 days in agar plates on basal medium deprived of N or supplemented with 1 mM NO₃⁻. The expression was quantified by RT-qPCR and normalized by *CLATHRIN*.

The data are the mean ± se from 3 independent experiments. P values were analyzed by Student's T Test and differences between genotypes are statistically significant at *P* < 0.005.

**Fig. 4: TAR2 expression in the roots is repressed in the absence of NO₃⁻ in a NRT1.1-dependent manner.**

Wild-type, *chl1-5* and *tar2-c* seedlings were grown for 12 days in vertical agar plates on basal medium deprived of N, or supplemented with either 1 mM NO₃⁻, or 0.5 mM L-glutamine.

(A) Relative accumulation of *TAR2* transcripts in roots. The data are the mean ± se of 2 replicates from 3 independent experiments. P values were analyzed by Student's T Test and differences between genotypes are statistically significant at **P** < 0.01.

(B) Histochemical staining of GUS activity in roots of plants expressing a *pTAR2::GUS* transgene in either wild type (Col) or *chl1-5* genetic background. Black bar represents 0.5 cm.

(C) Histochemical staining of GUS activity in the root stele and lateral root primordia of plants expressing a *pTAR2::GUS* transgene in either wild type (Col) or *chl1-5* genetic
background. The pictures shown display root portions with lateral root primordia at stages I-III. The pictures shown are representative of 10 primordia from >10 plants of two independent experiments. Black bar represents 0.5 cm.

**Fig. 5: Mutation of TAR2 represses DR5::GFP expression in LRPs and in the stele under NO₃⁻ supply.**

(A) DR5::GFP expression in mature root zone of plants with WT (left panel) or *tar2c* (right panel) genetic background. The seedlings were grown for 12 days on basal medium supplemented with 1mM NO₃⁻. White bars = 100 µm.

(B) DR5::GFP expression in lateral root primordia of plants with WT (upper panel) or *tar2c* (lower panel) genetic background. The seedlings were grown for 12 days on basal medium supplemented with 1mM NO₃⁻. The pictures display lateral root primordia at different developmental stages (from stage II to VI). The asterisk visualizes the location of the primordium. The pictures are representative of 10 plants of 2 independent experiments. White bars = 100 µm.

(C) Fluorescence quantification in lateral root primordia at different developmental stages of 12-d-old plants expressing DR5::GFP in WT or *tar2c* background. Plants were grown on basal medium supplemented with 1mM of NO₃⁻. Data represent mean ± se of 2 independent experiments (n= 10). P values were analyzed by Student's T Test and differences between genotypes are statistically significant at *, P < 0.05; **, P < 0.01) ***, P < 0.001.

**Fig. 6: Regulation of lateral root development by NO₃⁻ and NRT1.1 relies on the TAR2 auxin biosynthesis enzyme.**

Root system architecture of Col, *chl1-5*, *tar2c* and *chl1-5tar2c* plants grown for 12 days in vertical agar plates on a basal nutrient medium deprived of N or supplemented with either 1 mM NO₃⁻ or 0.5 mM Gln.

(A) Representative pictures of Col, *chl1-5*, *tar2c* and *tar2c*×*chl1-5* plants. White bar represents 0,5 cm.

(B) Mean length the primary root. Results (n=100) are representative of 5 independent experiments. P values were analyzed by Student's T Test and differences between mutant and wild-type genotypes are statistically significant at ***, P < 0.001 (t test).
(C) Mean density of visible lateral roots (> 0,5 mm). Results (n=100) are representative of 5 independent experiments. P values were analyzed by Student's T Test and differences between mutant and wild-type genotypes are statistically significant at ***, P < 0.001 (t test).

**Fig. 7: Schematic model of the integrated repressive role of NRT1.1 on lateral root development.**

NRT1.1 coordinately controls three separate auxin-related processes for repressing LRP development at low NO₃⁻ availability. First, NRT1.1 facilitates the basipetal transport of auxin out of the LRP (Krouk et al. 2010). Second NRT1.1 represses local auxin biosynthesis by downregulation the expression of the *TAR2* auxin biosynthetic gene, which reduces the acropetal flux of auxin to the LRP. These two actions are expected to lower auxin accumulation at the LRP apex that is required for normal LRP development and growth. Third, NRT1.1 maintains a mechanical resistance to the LRP outgrowth by repressing LAX3 in the tissues overlying the LRP, thereby preventing cell wall remodeling in these tissues. All three repressive effects of NRT1.1 are suppressed by high NO₃⁻ availability (>0.5 mM), leading to the local NO₃⁻-induced stimulation of LRP development and emergence.
<figure><img src="image_2.png" /><figcaption>Figure 1.</figcaption></figure>
<figure><img src="image_3.png" /><figcaption>Figure 2</figcaption></figure>
<figure><img src="image_4.png" /><figcaption>Figure 3</figcaption></figure>
<figure><img src="image_5.png" /><figcaption>Figure 4</figcaption></figure>
<figure><img src="image_6.png" /><figcaption>Figure 5</figcaption></figure>
<figure><img src="image_7.png" /><figcaption>Figure 6</figcaption></figure>
<figure><img src="image_8.png" /><figcaption>Figure 7</figcaption></figure>
<table><caption>Table 1. Primers used for qRT-PCR.</caption><thead><tr><th>Description</th><th>AGI</th><th>Primer</th><th>Sequence (5'-3')</th></tr></thead><tbody><tr><td rowspan="2">CLATHRIN</td><td rowspan="2">AT4G24550</td><td>Forward</td><td>AGCATACACTGCGTGCAAAG</td></tr><tr><td>Reverse</td><td>TCGCCTGTGTCACATATCTC</td></tr><tr><td rowspan="2">AUX1</td><td rowspan="2">AT2G38120</td><td>Forward</td><td>GAAGCCACCGTTCTTTATGC</td></tr><tr><td>Reverse</td><td>GCTGCAGCTGGTTTACATTG</td></tr><tr><td rowspan="2">LAX3</td><td rowspan="2">AT1G77690</td><td>Forward</td><td>TTGTGAGGAGGGCATTGGTA</td></tr><tr><td>Reverse</td><td>GGAGAGGCCACCGAGAGT</td></tr><tr><td rowspan="2">ABCB1</td><td rowspan="2">AT2G36910</td><td>Forward</td><td>CGACGGGAAAGACATAAGGA</td></tr><tr><td>Reverse</td><td>CTGGTAGCGCGGATATGAAT</td></tr><tr><td rowspan="2">ABCB4</td><td rowspan="2">AT2G47000</td><td>Forward</td><td>CGTGTCGTGCTTTCCCTTTCT</td></tr><tr><td>Reverse</td><td>ATCCAAAGGTGTTGCTGCTC</td></tr><tr><td rowspan="2">ABCB19</td><td rowspan="2">AT3G28860</td><td>Forward</td><td>CCAGCAAGAGGGTCGTAAAA</td></tr><tr><td>Reverse</td><td>GAGACGATCCGTGGAGACAT</td></tr><tr><td rowspan="2">PIN1</td><td rowspan="2">AT1G73590</td><td>Forward</td><td>TACTCCGAGACCTTCCAACTACG</td></tr><tr><td>Reverse</td><td>TCCACCGCCACCACTTCC</td></tr><tr><td rowspan="2">PIN2</td><td rowspan="2">AT5G57090</td><td>Forward</td><td>CCTCGCCGCACTCTTTCTTTGG</td></tr><tr><td>Reverse</td><td>CCGTACATCGCCCTAAGCAATGG</td></tr><tr><td rowspan="2">PIN3</td><td rowspan="2">AT1G70940</td><td>Forward</td><td>GAGGGAGAAGGAAGAAAGGGAAC</td></tr><tr><td>Reverse</td><td>CTTGGCTTGTAATGTTGGCATCAG</td></tr><tr><td rowspan="2">PIN4</td><td rowspan="2">AT2G01420</td><td>Forward</td><td>TTGTCTCTGATCAACCTCGAAA</td></tr><tr><td>Reverse</td><td>ATCAAGACCGCCGATATCAT</td></tr><tr><td rowspan="2">PIN7</td><td rowspan="2">AT1G23080</td><td>Forward</td><td>CGGCTGATATTGATAATGGTGTGG</td></tr><tr><td>Reverse</td><td>GCAATGCAGCTTGAACAATGG</td></tr><tr><td rowspan="2">LDB29</td><td rowspan="2">AT3G58190</td><td>Forward</td><td>GCACATTGGCATTAGGCTTT</td></tr><tr><td>Reverse</td><td>CTCAATCCCCACCTCAGCTA</td></tr><tr><td rowspan="2">TAR2</td><td rowspan="2">AT4G24670</td><td>Forward</td><td>CAGATTTGGCTTACTATTTGGCCACAG</td></tr><tr><td>Reverse</td><td>GTCTTTCACCAAAGCCCATCCCAATC</td></tr></tbody></table>