# Root energy availability controls the fast decrease in root nitrate uptake in response to light intensity decrease

Yifei Zhao, Thibaut Perez, Benoit Lacombe, Leo F M Marcelis

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Yifei Zhao, Thibaut Perez, Benoit Lacombe, Leo F M Marcelis. Root energy availability controls the fast decrease in root nitrate uptake in response to light intensity decrease. Plant Physiology and Biochemistry, 2025, 228, pp.110229. 10.1016/j.plaphy.2025.110229. hal-05162504

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# Root energy availability controls the fast decrease in root nitrate uptake in response to light intensity decrease

Yifei Zhao<sup>a, ●</sup>, Thibaut Perez<sup>b</sup>, Benoit Lacombe<sup>b, ●</sup>, Leo F.M. Marcelis<sup>a, *</sup>

## ARTICLE INFO

**Keywords:**

Nitrate uptake

Photosynthesis

Nitrate influx

Nitrate transporters

## ABSTRACT

Photosynthesis determines plant growth, influencing plant nitrogen demand, hence it is also crucial to the regulation of nitrate uptake. It remains unclear how fast plants adjust root nitrate uptake in response to change in photosynthesis. This research aimed to investigate the instantaneous response of root nitrate uptake to shoot photosynthesis. Arabidopsis grown hydroponically in a climate chamber under 175 µmol m⁻² s⁻¹ photosynthetically active radiation (PAR). On day 34, plants were transferred to three light intensities (75, 175, or 300 µmol m⁻² s⁻¹ PAR). Nitrate uptake, expression levels of four nitrate transporters, plant nitrogen (N) content, and plant mass were measured before transfer, 0.5 h, and 3 h after the transfer. CO₂ gas exchange was measured continuously. Photosynthesis rate immediately adjusted to new light intensity, reaching a steady state 20 min after the transfer. Nitrate influx and root sugar concentration both decreased in plants exposed to low-light intensity. Conversely, nitrate influx remained constant over the 3 h in plants transferred to high light, despite increased sugar concentration in the root. Only the expression level of NRT1.1 was upregulated in plants transferred to high light. Our findings highlight the importance of energy for nitrate uptake under low light condition. It also suggests that, within the initial 3 h following the light intensity transition, plants can maintain growth independent of nitrate uptake, likely through the assimilation of stored nitrogen. This demonstrates plants' ability to sustain growth without instance response in nitrate uptake and highlights the importance of energy availability in nitrate uptake.

## 1. Introduction

Plants face various and often unpredictable environmental challenges throughout their life cycles, requiring rapid and efficient adaptation strategies. Central to these adaptations is the coordination between root and shoot activities through both short- and long-distance signaling mechanisms. Nutrient uptake by roots is particularly crucial, enabling plants to acquire essential elements vital for growth and development. Nitrate uptake is a tightly regulated process influenced by several physiological factors. Previous studies have highlighted a correlation between photosynthesis and nitrate uptake, showing that light intensity stimulates nitrate transport (Delhon et al., 1995b; Lejay et al., 1999, 2008). For instance, nitrate uptake exhibits a diurnal rhythm, peaking during light periods and declining in darkness (Delhon et al., 1995a; Matt et al., 2001; Okuyama et al., 2015).

The relationship between light and nitrate uptake is complex, and

sugar is one of the main component that is involved in this regulation. External application of sugar to roots can sustain nitrate uptake levels even after plants are exposed to darkness, indicating that the effect of light on nitrate uptake is tightly related to the sugar level in plant (Delhon et al., 1995b; Lejay et al., 2003). Photosynthesis produces sugars that provide energy for the active transport of nutrients, including nitrate, into root cells. This transport process involves specific transport proteins in the root cell membranes, which move nitrate ions against their electrochemical gradient, indirectly using ATP energy (Britto and Kronzucker, 2006).

Additionally, sugars act as signaling molecules regulating nitrate uptake, directly modulating nitrate uptake, either directly at the root or indirectly by altering the plant's carbon status (Fichtner et al., 2021). This regulation primarily occurs at the molecular level, where the expression of nitrate transporters such as NRT1.1 and NRT2.1 is upregulated during light periods and downregulated in darkness, reflecting a

This article is part of a special issue entitled: Membrane transporters published in Plant Physiology and Biochemistry.

E-mail address: leo.marcelis@wur.nl (L.F.M. Marcelis).

<footnote><sup>a</sup> Horticulture and Product Physiology Group, Wageningen University and Research, P.O. Box 16 6700AA, Wageningen, the Netherlands</footnote>

<footnote><sup>b</sup> IPSiM, Université de Montpellier, CNRS, INRAE, Institut Agro, Montpellier Cedex, France</footnote>

<footnote>* Corresponding author.</footnote>
diurnal pattern (Lejay et al., 2003).

Furthermore, increased growth driven by photosynthesis elevates the plant's demand for nitrogen (N), potentially further altering the N status and C-N balance in different tissues. To maintain N homeostasis, plants adjust their uptake according to their needs, ensuring optimal growth and development (Crawford and Glass, 1998; Imsande and Touraine, 1994). This adjustment involves regulating nitrate transporters sensitive to the plant's nitrogen status. For example, the expression of the NRT2.1 transporter is influenced by the plant's overall nitrogen status (Girin et al., 2007; Glass et al., 2001; Lejay et al., 1999; Ruffel et al., 2011).

While the correlation between nitrate uptake and light intensity has been well-established, most studies have focused on prolonged exposure to different light intensities (Lejay et al., 2003; Sakuraba and Yanagisawa, 2018). However, in natural and agronomic environments, as well as in greenhouses, light intensity fluctuates throughout the day. Therefore, it is crucial not only to understand the general regulation of light on plant nitrate uptake but also to comprehend how plants adjust their nitrate uptake strategies in response to changing light conditions. Current knowledge suggests that photosynthesis responds to varying light intensities often within 10 min (Kaiser et al., 2018). However, root activities, particularly nitrate uptake, appear to respond more slowly to changes in light intensity, possibly due to the time needed for signals from the shoot to reach the roots (Fichtner et al., 2021).

In this paper, we quantify the dynamics of plant nitrate uptake in response to a step change in light intensity. We investigate whether the delay in response is due to the slow transport of sugar from the shoot to the root or if root nitrate uptake regulation is inherently slower. Additionally, we explore the implications of these differences in response times for the carbon-to-nitrogen status of plants.

## 2. Material and methods

### 2.1. Growth conditions and treatments

The study was conducted in a fully controlled climate chamber. Temperatures were maintained at 22 °C during the day and 20 °C during the night, with a photoperiod of 10 h. Relative humidity was kept at 70 % and CO2 concentration was ambient. Plant-level light intensity (PAR) was set at 175 µmol µmol m⁻² s⁻¹ using Phillips TL-D 58 W/40 lamps from the day of sowing. 34 days after sowing (DAS), 45 min after the light switched on in the climate chamber, plants were transferred from the initial light intensity of 175 µmol m⁻² s⁻¹ to different levels (75 and 300 µmol m⁻² s⁻¹, or maintained at 175 µmol m⁻² s⁻¹). The chamber was divided into three compartments, each receiving a specific light intensity treatment. Measurements were conducted at three time-points on the treatment day: just before, 30 min, and 3 h after the transfer to different light intensities. At each time point and under each light intensity treatment, plants from individual growth tanks were sampled for nitrogen (N), sugar, and gene expression analysis. These measurements were repeated over seven individual days, with treatments replicated each day. Gas exchange measurements were conducted on a single plant throughout the entire period for each light intensity treatment, repeated over 10 individual days with each treatment replicated daily.

### 2.2. Seed germination and plant growth

The seed germination was conducted following the protocol described by van Delden et al. (2020). On the 23rd days after sowing (DAS), seedlings were transplanted into growth tanks (30 × 20 × 22 cm, Euronormbak® reference RC-204) filled with 20 L of nutrient solution with a nitrate concentration of 1 mM (Table 1). Each growth tank accommodated four seedlings. The nutrient solution was aerated by an air pump and was covered with a polypropylene (PP) lid (30 × 20 cm, Euronormbak®). Considering that the N concentration in the nutrient solution remained relatively constant during the growth period (plants

<table><caption>**Table 1**

Nutrient solution composition.</caption><thead><tr><th colspan="2">Macronutrients (mM)</th></tr></thead><tbody><tr><td>N-NO<sub>3</sub></td><td>1</td></tr><tr><td>P-H<sub>2</sub>PO<sub>4</sub></td><td>0.5</td></tr><tr><td>K</td><td>3</td></tr><tr><td>Ca</td><td>2.5</td></tr><tr><td>Mg</td><td>1</td></tr><tr><td>S-SO<sub>4</sub></td><td>3</td></tr><tr><td>Cl</td><td>2.5</td></tr></tbody><thead><tr><th colspan="2">Micronutrients (mM)</th></tr></thead><tbody><tr><td>Fe</td><td>0.05</td></tr><tr><td>B</td><td>0.05</td></tr><tr><td>Cu</td><td>0.0005</td></tr><tr><td>Zn</td><td>0.01</td></tr><tr><td>Mn</td><td>0.005</td></tr><tr><td>Mo</td><td>0.0001</td></tr></tbody></table>

were grown in this system from 23rd DAS to 34th DAS), we did not add any water or nutrients during this period, except for adding citric acid to maintain pH at 5.8.

### 2.3. Nitrate influx, nitrogen content and nitrogen allocation

At 34 DAS, nitrate influx measurements were conducted following the methodology described by Delhon et al. (1995b). To begin, the roots were immersed in 70 ml of 0.1 mM CaSO₄ for 1 min to remove excess ¹⁴N from the root surface. Subsequently, the plant was transferred to a 50 ml labeling solution for 5 min. All macro- and micronutrient concentrations in this labeling solution were identical to those in the nutrient solution for plant growth, with the exception of isotopic ¹⁵N-nitrate being used in place of ¹⁴N-nitrate. Finally, the plant was transferred to a new container with 70 ml of 0.1 mM CaSO₄ for an additional minute. After these steps, the plant was harvested, and the shoot and root were separated to determine dry weight. The shoot and root samples underwent initial oven-drying (WTB binder 7200) at 40 °C for 24 h, followed by an additional 48 h at 70 °C until a consistent dry weight was achieved. The determination of the ¹⁵N/¹⁴N ratio, as well as the quantification of N content in shoot and root, were performed by Isotopes Quantifications platform (AQUI, IPSIM, France), using an Isoprime mass spectrometer coupled with a Pyrocube elemental analyzer (Elementar, Germany). Plant nitrogen allocation between shoot and root was calculated as the ratio of nitrogen content in the shoot to that in the root, with nitrogen content derived by multiplying nitrogen concentration (mg g⁻¹ dry weight) by the respective organ's dry weight (g).

### 2.4. Soluble sugars and starch analysis

Shoot or root tissues were initially freeze-dried (Edwards 4K Mod-ulyo FreezeDryer) and ground to a fine powder using a ball mill. To extract soluble sugars, 3 mg of powdered tissue was mixed with 1 ml of 80 % ethanol. The mixture was vortexed (Scientific Industries, Vortex genie-2) then transferred to an ultrasonic bath (Branson 2800) for 10 min. After another round of vortexing, the samples were placed in a shaking water bath (GFL 1083) at 80 °C for 20 min, followed by a 3rd round of vortex. The samples were centrifuged at 25000 rcf for 5 min (Sorvall Legend Micro 21R (ThermoScientific)). For further sugar determination, 0.8 ml of the supernatant was transferred to a new 1.5 ml Eppendorf tube. Simultaneously, the pellet with the remaining supernatant was preserved for starch determination.

The 0.8 ml supernatant for sugar determination was then dried using a Speedvac vacuum centrifuge (Savant SPD2010 SpeedVac) at 45 °C and 5.1 Torr for 2 h. Subsequently, 1 ml Milli-Q water was added to each dried sample, and the tubes were vortexed. The samples underwent another round of ultrasonication for 10 min, followed by vortexing and a final centrifugation step of 10 min at 25,000 rcf. Finally, the samples were diluted tenfold to determine the concentration of sucrose, glucose,
and fructose using a High-Performance Anion Exchange Chromatography with Pulsed Amperometric Detection (HPAEC-PAD; Dionex ICS5000, Thermo Fisher Scientific, Waltham, MA, United States), equipped with a CarboPac PA1 column (250 mm × 2 mm, Thermo Fisher Scientific). The elution was carried out using 100 mM NaOH at a flow rate of 0.25 mL/min at 25 °C.

For starch determination, the stored pelleted was initially rinsed three times with 80 % ethanol to remove the remaining sugar. Subsequently, the pellet was vacuum-dried at 50 °C and 5.1 mbar for 20 min using a centrifuge. Following drying, the pellet was resuspended in 0.8 mL of 0.5 g L⁻¹ amyloglucosidase (Sigma 10115) in 50 mM citrate buffer (pH 4.6), followed by a 15-min incubation at 60 °C. After centrifugation at 21,100 RCF for 5 min, glucose quantification was performed using HPEAC-PAD as described above, with starch data expressed based on root dry weight as mg g⁻¹ DW after 10–20 times dilution with Milli-Q water.

## 2.5. Photosynthetic gas exchange

At 34 DAS, plants designated for photosynthetic gas exchange measurements were first transferred to the corresponding treatment cabinet with corresponding light intensity before the light in the climate chamber was switched on. This precaution was taken to avoid potential physical damage during movement. Measurements were conducted using the LI-6800 photosynthesis system (LI-COR Biosciences, Lincoln, NE, USA) equipped with a leaf chamber fluorometer (LI-COR part no. 6400–40, enclosed leaf area: 2 cm²).

Environmental conditions in the leaf chamber, except for light intensity, were maintained to match those in the climate chamber (400 ppm CO₂, 22 °C Tₐir, and RHₐcham 70 %). Gas exchange measurement began 15 min after the light was switched on in the climate chamber. The initial light intensity in the leaf chamber was set at 175 µmol m⁻² s⁻¹ with a light spectrum of 90 % red and 10 % blue, mimicking the initial light conditions of all treatments at the beginning of the day. Subsequently, the light intensity in leaf chambers was adjusted to 75. 175, or 300 µmol m⁻² s⁻¹, corresponding to the specific treatment of the plants. Measurements were performed on fully expanded and unshaded leaves. Net photosynthesis rate, transpiration rate, and stomatal conductance was continuously recorded up to 3 h following the light intensity adjustment.

## 2.6. Gene expression analysis

Quantitative reverse transcription polymerase chain reaction (RT-qPCR) was used to determine the expression levels of genes involved in nitrate transport (NRT1.1, NRT1.2, NRT2.1, NRT2.2). RNA was extracted from fine powders of pooled fresh frozen root samples using the CiAR method (Oñate-Sánchez and Verdonk, 2021). The purity and concentration of the extracted RNA were measured with a Nanodrop™ One Microvolume UV–Vis Spectrophotometer (Thermo Scientific™) and verified by electrophoresis on a 1.5 % w/v agarose gel with 200 ng of RNA. Following RNA extraction, DNAse treatment and cDNA synthesis were performed according to the manufacturer's instructions. The resulting cDNA was diluted five-fold for subsequent use.

Gene expression was analyzed in a 10 µL reaction mixture, which included 5 µL of SYBR-green master mix (Bio-rad, USA), 0.5 µL each of forward and reverse primers (10 µM), 3 µL of Milli-Q water, and 1 µL of cDNA. The thermal cycling was conducted using a CFX96 thermal cycler (Bio-rad). Specific primers for the target genes were designed using Primer3Plus. To normalize gene expression levels, ACT2 and APT1 were used as internal reference genes. Detailed information and primer sequences for the target genes are provided in the Supplementary material (Supplementary Table 1). Each qPCR analysis was performed with three biological replicates, each containing three technical replicates. The efficiency of the qRT-PCR reactions and R values were assessed using the LinRegPCR program (Ruijter et al., 2009). Ct values were recorded

under default conditions. Relative gene expression (RGE) levels were calculated using the 2⁻⁰ΔCt method (Livak and Schmittgen, 2001).

## 2.7. Statistics

This study used a randomized complete block design with seven blocks (n = 7). Within each block, one growth tank represented one true replicate for one treatment, and each tank contained six plants that acted as pseudo replicates. For growth parameters determinations, influx measurements, and N content analysis, one statistical replicate is one plant per container. Soluble sugars and starch analysis and gene expression analysis were based on a pooled sample from the remaining five plants from the same container.

All data was analyzed in R version 4.2.1. Linear mixed effects regression models, i.e. the lmer function from the lme4 package version 1.1–30, were used to assess for differences in the least square (LS) means of the response variables. For each response variable, the light intensity treatment was taken as main effect with blocks as random effects. Assessment for significant differences (P < 0.05) was performed using Tukey adjusted least square means (emmeans version 1.7.5). The residuals of the lmer models were tested for normality (Shapiro–Wilkinson test and QQ-plots inspection) and homogeneity (bartlett test and histogram inspection).

## 3. Results

After 0.5 h, there were no statistically significant differences in nitrate influx among the three light intensities, although a trend emerged where lower light corresponded to slightly reduced influx (Fig. 1). By 3 h, these differences became pronounced: at high light intensity (300 µmol m⁻² s⁻¹), nitrate influx remained stable over time; at medium

<figure><img src="image_3.png" /><figcaption>Fig. 1. Nitrate influx in root of 34-day-old arabidopsis Col-0 responding to the change of light intensity over time. Each data point represents the mean of 7 replicate plants. Error bars represent the standard error of the mean. Means that share the same letter, are not significantly different (P < 0.05, Tukey's test).</figcaption></figure>
intensity (175 µmol m⁻² s⁻¹), it declined slightly but not significantly; and at low intensity (75 µmol m⁻² s⁻¹), it decreased significantly. Consequently, after 3 h, nitrate influx levels clearly reflected the applied light intensities, with the highest light condition maintaining the greatest influx and the lowest light condition showing the most pronounced decline.

Nitrogen content in shoot decreased from hour 0 to hour 3 when plants were transferred to high light intensity (Fig. 2 a). This reduction trend was observed in plants under medium and high light intensity throughout the 3 h but was not observed in plants transferred to low light intensity. In contrast, nitrogen content in root remained constant among all light levels during the whole period (Fig. 2 b).

Across all light intensities, plant dry weight showed a non-
statistically significant trend of increase over the 3 h following the
transfer to different light intensities, with the least increase observed
under low light intensity (Fig. 3 a). The nitrogen content per plant and
nitrogen allocation between shoot and root remained unaffected by light
intensities throughout the 3-h duration (Fig. 3 b c).

Photosynthesis rates instantly responded to light treatments and reached new steady states within 15 min after the transfer, with high light resulting in an increase, low light resulting in a decrease, and medium light maintaining a relatively constant photosynthesis rate (Fig. 4 a). Afterwards, photosynthesis rates remained at a relatively stable level in each light treatment. Upon decrease of light intensity to 75 µmol m² s⁻¹, transpiration rate and stomatal conductance declined for about 20 min, after which they increased slightly (Fig. 4 b c). Upon increase of light intensity to 300 µmol m² s⁻¹, both parameters gradually increased to slightly higher values than at the medium light intensity (175 µmol m² s⁻¹).

High light intensity resulted in higher soluble sugar content in shoot (Fig. 5 a). The high light intensity treatment exhibited an increase in shoot soluble sugar content throughout the entire period post-transfer. For the medium light treatment where light intensity remained constant, the total soluble sugar content remained rather constant as well, while the transfer of plants to the low light intensity led to a decrease in sugar content within 0.5 h after the transfer, persisting until 3 h post-transfer.

The starch concentration in shoot increased linearly over time at all intensities (Fig. 5 c). High light intensity resulted in the highest starch content, while low light intensity resulted in the lowest content.

Higher nitrate influx correlated with higher root total soluble sugars

(R² = 0.71, p < 0.05) in plants across all light intensities, following a polynomial trend (Fig. 6).

To investigate the immediate response of nitrate uptake activity to varying light intensities, we examined the relative gene expression (RGE) levels of four pivotal nitrate transporters in root tissues at different time points after the transfer to different light intensities.

Half an hour after the transfer, NRT1.1 showed an increase (2-fold) in RGE in plants transferred to high light and a decrease in plants transferred to low light (Fig. 7 a). However, this response was no longer observed at the 3-h mark after the transfer.

At 0.5 h after the transfer, there seemed to be an increase in RGE of NRT2.1 under all light conditions, although not significantly(Fig. 7 b). At 3 h, a notable increase in RGE was observed in plants transferred to high light.

The RGE of NRT1.2 remained constant across all light intensities over time, except for plants transferred to high light intensity, where a significant increase in RGE of NRT1.2 was observed from 0.5 to 3 h after the transfer (Fig. 7 c). Similarly, the RGE of NRT2.2 was not affected by light intensity (Fig. 7 d). However, a general trend of increase in the RGE of NRT2.2 was observed over time across all light intensities.

4. Discussion

4.1. Role of energy availability in nitrate uptake

In this study, we quantitatively characterized the dynamics of plants nitrate uptake in response to changes in light intensity. When plants were transferred to low light conditions, nitrate influx significantly decreased within half an hour (Fig. 1). This swift modulation in nitrate uptake may correlate with the constrained energy availability resulting from reduced photosynthesis (Fig. 4). A possible explanation is that photosynthetically derived sugars, such as sucrose, activate plasma membrane H + ATPase (Kinoshita et al., 2023), thereby influencing nitrate uptake rates. As photosynthesis rates declined, there was a concurrent decrease in sugar content in both shoot and root tissues, within 0.5 h in the shoot and 3 h in the root (Fig. 5). The slower decrease in root sugar content compared to the shoot is likely due to the predominant synthesis of sugars in the shoot via photosynthesis. Given the energy-demanding nature of nitrate influx (Britto and Kronzucker, 2006), the decline in root sugar availability under low light intensity likely limits the energy resources necessary for sustaining nitrate uptake,

<figure><img src="image_4.png" /><figcaption>**Fig. 2.** Nitrogen content per gram shoot dry weight (SDW) (a) and per gram root dry weight (RDW) (b) of 34-day-old arabidopsis Col-0 responding to the change of light intensity over time. Each data point represents the mean of 7 replicate plants. Means that share the same letter, are not significantly different (P < 0.05, Tukey's test).</figcaption></figure>
<figure><img src="image_5.png" /><figcaption>**Fig. 3.** Plant dry weight (a), nitrogen content per plant (b), and shoot-to-root nitrogen content ratio (c) of 34-day-old arabidopsis Col-0 responding to the change of light intensity over time. Each data point represents the mean of 7 replicate plants. Error bars represent the standard error of the mean (when larger than symbol size). Means that share the same letter, are not significantly different (P < 0.05, Tukey's test).</figcaption></figure>

contributing to the observed reduction in nitrate influx rates. This hypothesis is supported by the correlation between nitrate influx and root sugar content (Fig. 6). Changes in stomatal conductance and transpiration rate under different light intensities (Fig. 4 b c) may also influence nitrate uptake, likely through effects on xylem flow (Orieux et al., 2018). While not the primary focus of this study, these factors may contribute to the reduced uptake observed under low light.

When light intensity increased from 175 to 300 µmol m⁻² s⁻¹, within the treatment itself, there was no statistically significant increase in nitrate influx over the 3-h period following the transfer, despite a notable increase in root sugar content (Figs. 1 and 5). Although high light successfully increases root energy availability, indicated by a 20 % increase in root soluble sugars, this surplus energy did not lead to a corresponding rise in nitrate influx. Previous studies have shown that nitrate uptake can be induced by light or sugar, but often conducted experiments under light/dark switch conditions (Delhon et al., 1995b; Lejay et al., 2008). In such scenarios, plants experience drastic changes in light intensity, leading to more pronounced changes in root sugar content and, consequently, energy availability for nitrate influx. However, in our study, the change in light intensity is relatively mild; both high and low light treatments started from an initial light intensity of 175 µmol m⁻² s⁻¹, which is suitable for arabidopsis cultivation. Therefore, the initial root sugar content was relatively high, whilst the changes in root sugar content over time upon the changes in light intensity were relatively small.

We propose that the relationship between energy and nitrate influx most likely follows a saturation response. Initially, increasing energy levels lead to a proportional increase in nitrate influx, as observed in previous studies (Delhon et al., 1995a, 1995b; Lejay et al., 2008). However, beyond a certain energy threshold, further increases do not result in additional nitrate influx, as demonstrated by the high light treatment in this study (Fig. 6). This suggests that nitrate uptake is not solely driven by available energy but is also regulated by complex feedback mechanisms involving nitrogen and carbon metabolism. For example, AtNRT2.1 has been shown to integrate signals from both nitrogen and carbon status, limiting nitrate uptake under high-energy conditions to match metabolic demand (Lejay et al., 2003). Liang et al. (2022) also provided evidence of this regulatory complexity, showing that nitrogen uptake and allocation between roots and shoots are dynamically adjusted to balance tissue-specific metabolic needs.

## 4.2. Response of expressions of nitrate transporters to light intensity

Nitrate transporters, which facilitate the movement of nitrate ions (NO₃⁻) across the cell membrane, play an important role in regulating nitrate influx. In this study, we investigated the relative gene expression (RGE) levels of four nitrate transporters expressed in root tissues to elucidate their response to changes in light intensity. These transporters were selected because they represent two nitrate uptake mechanisms: high-affinity transport systems (HATS) and low-affinity transport systems (LATS). NRT2.1 and NRT2.2 contribute to HATS, facilitating rapid nitrate uptake when external nitrate concentration is low. NRT1.2, on the other hand, is involved in LATS, which mediates nitrate uptake when external nitrate concentration is high. NRT1.1 is unique as a dual-affinity nitrate transporter, participating in nitrate influx under both low and high nitrate concentration conditions. We chose to study nitrate influx and nitrate transporter expression levels when arabidopsis plants were grown at an external nitrate concentration of 1 mM because, as at this concentration the high-affinity system is saturated, and the low-affinity system begins to play a more significant role in nitrate uptake (Lyzenga et al., 2023). Our aim was to elucidate how these transporters respond to varying light intensities, providing insight into the regulation of nitrate uptake in plants under different environmental conditions.

For plants transferred to high light, we observed that, among the four nitrate transporters tested, only NRT1.1 exhibited increased expression level in plants 0.5 h after the transfer to high light intensity (Fig. 7). A correlation between the expression of NRT1.1 and NRT2.1 seemed to be consistent across different datasets or experimental conditions (reviewed by Ruffel et al., 2014). Although the direct relationship between NRT1.1 and plant carbon status is not explicitly detailed, plant C provision seemed to be involved in the regulation of NRT2.1 by influencing NRT1.1 (Muños et al., 2004). This suggests that NRT1.1 may play a primary role in adapting nitrate transport in response to the plant's C status compared to other nitrate transporters.

Previous research has demonstrated that nitrate transporters such as NRT1.1 and NRT2.1 are regulated by sugar and light (Miller et al., 2007; van Gelderen, 2021). However, in our study, taking the low light treatment as an example, none of the four nitrate transporters showed a significant decrease in expression levels 0.5 and 3 h after the transfer, despite a 14 % and 34 % reduction in nitrate influx rate observed in plants 0.5 and 3 h after being transferred to the low light, respectively (Figs. 1 and 7). This discrepancy suggests that the observed reduction in nitrate influx within the first 3 h of low light conditions is not primarily due to the downregulation of nitrate transporter expression. The
<figure><img src="image_6.png" /></figure>

<figure><img src="image_7.png" /></figure>

<figure><img src="image_8.png" /><figcaption>**Fig. 4.** Time course of net photosyntehsis rate (a), transpiration rate (b), and stomatal conductance (c) of 34-day-old arabidopsis Col-0 responding to the change of light intensity. Light intensities include 75, 175, and 300 µmol m⁻² s⁻¹ PAR, with distinct colors indicating each level. Initially, plants were exposed to a constant light intensity of 175 µmol m⁻² s⁻¹ PAR before being transferred to the specified light intensities. The vertical black line indicates the moment when plants were subjected to a step change in light intensity. Each data point represents the mean of 11–12 replicate plants.</figcaption></figure>

regulation of the activity of the transporters at the plasma membrane may still be involved via post-translational regulation (Chaput et al., 2023; Ho et al., 2009; Léran et al., 2015).

One reason for the lack of significant changes in expression levels may be the extent of the change in light intensity. Unlike some studies that transition plants from light to dark (Lejay et al., 1999, 2003, 2008), our study only adjusted light intensity from moderate to either low or high levels. This less drastic change may not be sufficient to trigger significant changes in transporter expression. For example, in studies where plants were transferred from dark to very high light intensity (800 µmol m⁻² s⁻¹) no difference in the expression level of NRT2.1 was observed until 4 h after the treatment (Ruffel et al., 2021). This indicates that both the extent and duration of light intensity change are crucial

factors in determining the expression response of nitrate transporters.

These results collectively indicate that factors other than changes in nitrate transporter expression levels are primarily responsible for modulating nitrate transport at the root level under varying light intensities, at least at the first 3 h when plants were subjected to change in light intensity. We conclude that while light intensity may influence the nitrate transport capacity of roots, its short-term effect on nitrate influx is not mediated solely through the regulation of nitrate transporter expression. While we hypothesize a regulatory role for NRT1.1 in adjusting nitrate uptake in response to light intensity, this conclusion is based on correlative data. Future studies should test this hypothesis using NRT1.1 mutant lines to verify its specific contribution to nitrate transport modulation under dynamic light conditions. Root-level factors, such as energy availability and related metabolic signals, likely play a predominant role in modulating nitrate transport dynamics under different light conditions (Lillo, 2004). This understanding aligns with previous research suggesting that root energy status and associated signaling pathways are critical determinants of nitrate uptake behavior (Delhon et al., 1995a; Lejay et al., 1999).

### 4.3. Did plants take what they needed?

To assess whether plants are meeting their N requirements for growth under varying light intensities, it is essential to consider the dynamic nature of plant N demand, which is influenced by both internal and external factors. Plants take up nitrate based on their N demand when there is adequate energy in the roots and available N in the rhizosphere (Imsande and Touraine, 1994). In this study, we hypothesized that changes in light intensity would primarily impact plant N demand, with plant growth rate being a key internal factor correlated with light intensity. Hence, when evaluating the effects of changing light intensity, the central question is whether plants can acquire sufficient N to support their growth. Both the supply (uptake) and demand for N may change in response to alterations in light intensity.

Plant dry weight tended to increase, though this effect was not statistically significant, within the initial 3 h of high-light treatment (Fig. 3). This increase was accompanied by a notable reduction in shoot nitrogen content (mg g⁻¹) in plants exposed to high and medium light intensities (Fig. 2). This reduction in nitrogen content may reflect enhanced nitrogen assimilation under high light-intensity conditions (Fu et al., 2017). Despite this potential plant growth, there is no corresponding increase in nitrogen acquisition in terms of nitrogen content (mg plant⁻¹) (Fig. 3). We expected plant demand to increase linearly with carbon accumulation. Hence, we hypothesize that plants subjected to high light experienced an elevated demand for nitrogen, outpacing the capacity for nitrate uptake within the 3-h timeframe. Consequently, nitrogen reserves within the plant were assimilated to sustain growth, resulting in a reduction of N per unit of shoot dry weight, illustrating a strategic reallocation of resources. Nitrogen allocation between shoot and root remained unchanged across treatments during the observation period (Fig. 3 c), suggesting that short-term light responses did not involve adjustments in N partitioning.

Conversely, even in conditions of reduced nitrate influx, such as the observed 34 % decline in plants transferred to low light, growth persisted. The observed reduction in nitrate influx appeared to still meet the diminished nitrogen demand, thus obviating the need for additional nitrogen uptake to support growth. Plants under low light conditions showed no noticeable change in N content despite the decrease in nitrate influx. However, it is important to note that plants are unlikely to be in a steady state within the 3-h duration of this study. The observed reduction in shoot nitrogen content indicates a mismatch between nitrate influx and increased plant growth in plants transferred to high light intensity. Furthermore, it suggests that the demand for nitrogen by the plants is still expected to persist given the ongoing accumulation of biomass and corresponding nitrogen requirements.

Two outcomes can be anticipated after the 3-h observation window:
<figure><img src="image_9.png" /><figcaption>**Fig. 5.** Total soluble sugars content per gram shoot dry weight (SDW) (a) and per gram root dry weight (RDW) (b), and starch content per gram shoot dry weight (SDW) (c) of 34-day-old arabidopsis Col-0 in response upon change of light intensity at time 0.0 from 175 to 75, 175, and 300 µmol m⁻² s⁻¹ PAR. Each data point represents the mean of 7 replicates, with each replicate consisting of pooled samples from 5 plants. Error bars represent the standard error of the mean. Means that share the same letter are not significantly different (P < 0.05, Tukey's test).</figcaption></figure>

<figure><img src="image_10.png" /><figcaption>**Fig. 6.** Nitrate influx plotted against total soluble sugars content in the root of 34-day-old Arabidopsis Col-0 responding to the change of light intensity over time. The scattered points represent the average of each treatment level, with different colors corresponding to light intensities of 75, 175, and 300 µmol m⁻² s⁻¹ PAR. Different symbols represent the value measured before, 0.5 h, and 3 h after the transfer. The light green dashed line represents the polynomial regression between the means of nitrate influx and root total soluble sugars under all light intensities at all time points.</figcaption></figure>

First, plant N demand may continue to rise as growth proceeds, potentially leading to a scenario where shoot N content (mg g⁻¹) drops below a critical threshold, limiting further growth. Second, the induction of nitrate transporters by light intensity may occur, resulting in the upregulation of nitrate influx to meet the increasing plant N demand. This regulatory response could potentially mitigate further reductions in shoot nitrogen content, allowing growth to proceed without sacrificing nitrogen reserves. Interestingly, root N content (mg g⁻¹) was unaffected by light intensity throughout the treatment period, while shoot N content decreased over time in plants transferred to medium and high light

(Figs. 1 and 2). This indicates distinct tissue-specific strategies for N regulation, with shoots showing higher N demand under high light that is not immediately balanced by nitrate uptake or assimilation.

Under low light, reduced photosynthesis limits the energy available for nitrate transport and assimilation. Lower ATP and reducing power (NADH/NADPH) constrain nitrate reductase (NR) activity, leading to nitrate accumulation in vacuoles or the cytosol instead of conversion to amino acids or proteins (Borbély et al., 2022; Huarancca Reyes et al., 2018; Jonassen et al., 2008). This nitrate buildup likely triggers feedback inhibition, downregulating root nitrate transporter expression via shoot-derived signals such as reduced cytokinin production (Andrews and Raven, 2022; Jonassen et al., 2008). As a result, nitrate uptake decreases, nitrogen assimilation slows, and growth is suppressed under low light.

In contrast, high light (300 µmol m⁻² s⁻¹) stimulates photosynthesis, enhancing NR activity in shoots and supporting rapid nitrate reduction. Nitrate reduction occurs predominantly in shoots, where energy is readily available (Kalcsits and Guy, 2013; Tang et al., 2012). This ammonium is quickly assimilated into amino acids and proteins via the GS/GOGAT pathway (Andrews and Raven, 2022). However, nitrate uptake does not significantly increase within 3 h, likely due to a lag in nitrate transporter induction or reliance on tissue nitrate reserves (Varala et al., 2018). Nitrogen assimilation supports accelerated growth, resulting in a dilution effect on shoot N concentration (mg g⁻¹) despite increased total nitrogen content (Luo et al., 2022). Over time, depletion of shoot nitrate pools may reduce feedback inhibition, enabling upregulation of nitrate transporters and improved nitrate uptake to meet rising N demand.

These findings highlight the dynamic interaction between light intensity, nitrate reduction, and nitrogen uptake. Under low light, reduced photosynthesis limits assimilation, suppresses nitrate uptake, and slows growth. High light enhances nitrogen assimilation and growth, but nitrate uptake lags, causing a temporary imbalance. Future studies should extend observation periods and include measurements of cytosolic nitrate pools and NR activity to better elucidate these regulatory mechanisms and improve our understanding of nitrogen homeostasis under varying light conditions.

## 5. Conclusion

Our study aimed to quantify the dynamics of how rapidly plants adjust their nitrate uptake strategy in response to fluctuations in light intensity. Our findings suggest that under low light conditions, nitrate
<figure><img src="image_11.png" /><figcaption>Fig. 7. Relative gene expression (RGE) of Nitrate transporter genes in the roots of 34-day-old arabidopsis Col-0 in response upon change of light intensity at time 0.0 from 175 to 75, 175, and 300 µmol m⁻² s⁻¹ PAR. Initially, plants were exposed to a constant light intensity of 175 µmol m⁻² s⁻¹ PAR before being transferred to the specified light intensities. Bar graphs represent the relative expression level of NRT1.1 (a), NRT2.1(b), NRT1.2 (c), and NRT2.2 (d) of all treatments relative to the control treatment which is before the transfer. All expression levels were measured in the roots using real-time quantitative PCR and normalized to two reference genes (APT1 and ACT2). Data are normalized means (±SE) obtained from 3 replicates constituted of a pool of 5 plants each. Means that share the same letter are not significantly different (P < 0.05, Tukey's test).</figcaption></figure>

influx is constrained by limited energy availability at the root. Surprisingly, after changing to high light conditions, nitrate influx remained unchanged, despite increased energy availability at the root, as suggested by increased sugar concentration. This highlights plants' ability to adapt photosynthesis, and likely growth, despite the lack of instant response in nitrate uptake.

### CRediT authorship contribution statement

**Yifei Zhao:** Writing – original draft, Visualization, Validation, Methodology, Investigation, Formal analysis, Data curation, Conceptualization. **Thibaut Perez:** Writing – review & editing, Methodology, Investigation. **Benoit Lacombe:** Writing – review & editing, Validation, Methodology, Conceptualization. **Leo F.M. Marcelis:** Writing – review & editing, Supervision, Resources, Project administration, Methodology, Conceptualization.

### Declaration of competing interest

The authors declare that they have no known competing financial interests or personal relationships that could have appeared to influence the work reported in this paper.

### Acknowledgement

We would like to thank Sarah Berman, Silvère Vialet-Chabrand, and Baltasar Zepeda for their valuable support and constructive feedback on this paper. We are also grateful to Gerrit Stunnenberg and David Brink of Unifarm for their help with the work in the climate chamber, and to Kjell Sneeuw for his significant assistance in the experiment.

### Appendix A. Supplementary data

Supplementary data to this article can be found online at https://doi.org/10.1016/j.plaphy.2025.110229.

### Data availability

Data will be made available on request.

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