Symbiotic nitrogen fixation suppresses root nitrate uptake in <i>Medicago truncatula</i> under nitrate limitation

While most plants use nitrate as the main nitrogen source (Gent & Forde, 2017), legumes such as Medicago truncatula can also acquire nitrogen by forming an endosymbiosis with N2-fixing bacteria called rhizobia, a process referred to as nodulation. In this endosymbiosis, the rhizobia are taken up into nodule cells which provide the low-oxygen environment required for N2 fixation (Layzell & Hunt, 1990). This energy-intensive process requires a constant supply of carbon from the host to produce ammonia that is directly assimilated in the nodule and then exported to the rest of the plant as amides or ureides, depending on the legume species (Ta et al., 1986; King & Purcell, 2005; Lu et al., 2022). To optimize growth and productivity, legumes need to maintain a balance between nitrogen gains and carbon losses during nodulation (Reid et al., 2011; Nishida & Suzaki, 2018); and this is why under nitrate-sufficient conditions, nodule formation and N2 fixation are suppressed (Chaulagain & Frugoli, 2021). Recently, it was shown in Lotus japonicus and Medicago truncatula that nitrate suppression of nodulation depends on NRT2.1s and the NIN-like protein (NLP) transcription factors that control their expression (Misawa et al., 2022; Luo et al., 2023). The suppression of nodulation by high nitrate levels presumably reflects the higher cost of symbiotic N2 fixation over direct N uptake from the soil (Zhao et al., 2024). However, how these two systems interact under low nitrate levels remains unknown. To investigate this, we examined the effect of nodulation on root nitrate uptake in M. truncatula. Experiments using labeled nitrogen (15NO3−) revealed a significant reduction of 15N content in both denodulated roots (i.e. root sections with nodules removed) and shoots of nodulated plants compared to the uninoculated controls at 3 wk postinoculation (wpi) (Fig. 1a). Direct nitrate uptake in plants occurs through high- and low-affinity transport systems which allow plants to adapt to varying soil nitrate levels (Crawford & Glass, 1998). Nitrate uptake from soil is mainly mediated by nitrate transporter 2 (NRT2) /nitrate assimilation related-protein 2 (NAR2) complexes and nitrate transporter 1 (NRT1) / peptide transporter (PTR) family (NPF) members (Gojon et al., 2011; Krapp et al., 2014; Noguero & Lacombe, 2016). We next examined whether NRT2.1, previously identified as a key nitrate uptake transporter in M. truncatula (Luo et al., 2023), might contribute to this process. We found that under low (0.2 and 0.5 mM) nitrate concentrations that are conducive for nodulation, the expression of MtNRT2.1 was significantly downregulated in denodulated roots compared to noninoculated controls 3 wpi with rhizobia (Fig. 1b). By contrast, for plants nodulated under high (5 mM) nitrate, no significant difference was observed in the 15N content of the shoots or roots, or in the levels of NRT2.1 expression in their denodulated roots, relative to uninoculated controls (Supporting Information Fig. S1a,b). Moreover, we found that this repression did not occur at earlier nodulation time points (Fig. S2). We then tested whether NRT2.1 is important for nitrate uptake under 0.2 mM KNO3. This revealed that the nrt2.1 mutant shoots and roots absorbed significantly less 15KNO3 than in the wild type (WT) (Fig. 1c), similar to what was reported for the mutant when grown under 0.5 and 5 mM KNO3 (Luo et al., 2023). These findings indicate that nodulation negatively impacts root nitrate uptake by downregulating MtNRT2.1 expression specifically under nitrate limiting conditions. To gain an insight into how this phenomenon is regulated, we performed RNA-seq analysis on denodulated roots and roots of noninoculated plants grown under 0.2 mM KNO3. A total of 5105 genes were upregulated, and 3540 genes downregulated in roots of nodulated plants (denodulated roots) compared to control roots. The full list of differentially expressed genes (DEGs) is provided in Table S1. Principal component analysis (PCA) of the RNA-seq results revealed a clear distinction between noninoculated and rhizobia-inoculated root samples for PC1, which explained 61% of the variation (Fig. S3a). A Kyoto Encyclopedia of Genes and Genomes (KEGG) term enrichment analysis revealed that denodulated roots have increased expression of genes related to ‘TCA cycle’, ‘glycolysis/gluconeogenesis’, ‘pyruvate metabolism’, ‘fructose and mannose metabolism’, and ‘starch and sucrose metabolism’, suggesting they have strongly enhanced sugar metabolism, while genes related to ‘nitrogen metabolism’ and ‘circadian rhythm’ were decreased (Fig. S3b). In addition, the results confirmed the repression of NRT2.1 in denodulated roots and revealed similar suppression of NRT2.2 and NAR2.1, which encode components of the high-affinity nitrate uptake complex, NLP4 which is required for nitrate suppression of nodulation, and its homolog NLP5 (Marchive et al., 2013; Xiao et al., 2021; Luo et al., 2023) (Fig. 1d). The ‘nitrogen metabolism’ genes that were downregulated also included the gene encoding the main root isoform of nitrate reductase, NR2, and two glutamate synthase genes, suggesting that nitrate assimilation is reduced in denodulated roots. Notably, members of two transcription factor families, Lateral Organ Boundaries Domain (LBD) proteins and Nitrate-Inducible GARP-type Transcriptional Repressor 1s (NIGT1s), thought to be involved in the repression of NRT2.1 expression in Arabidopsis thaliana and rice (Rubin et al., 2009; Yang et al., 2016; Kiba et al., 2018; Maeda et al., 2018; Zhu et al., 2022), were upregulated in roots of nodulated plants (Fig. 1d, see phylogenetic relationships in Figs S4, S5), indicating their possible role in regulating NRT2.1 expression. While NIGTs were reported to respond specifically to nitrate (Kiba et al., 2018; Maeda et al., 2018), LBDs have been reported to respond to different forms of fixed nitrogen, including ammonium and glutamine in A. thaliana and rice (Rubin et al., 2009; Zhu et al., 2022). To test whether ammonia could affect NRT2.1 expression, we evaluated its transcript levels 6 h after provision with (NH4)2SO4 in the presence of low concentrations of nitrate. The expression of NRT2.1 was induced by nitrate, and was similarly induced by a combination of ammonium and nitrate, relative to the nitrate-starved controls, and no induction was seen with ammonium (Fig. S6). This is similar to A. thaliana, where ammonium did not downregulate NRT2.1 expression under lower nitrate levels (Nazoa et al., 2003; Krouk et al., 2006), suggesting that ammonia produced in nodules may not be directly involved in the suppression of NRT2.1. We then investigated glutamine, which was shown to inhibit the expression of NRT2s in Nicotiana plumbaginifolia, A. thaliana, and Hordeum vulgare (Quesada et al., 1997; Krapp et al., 1998; Vidmar et al., 2000) and is a major export product of nodules. We first tested glutamine's effect on nonsymbiotic roots. Consistent with findings from other species, we found that the presence of glutamine strongly reduced the induction of MtNRT2.1 by nitrate (Fig. 1e). Furthermore, the expression of MtNIGT1.1 and MtNIGT1.2 was enhanced both by nitrate and by glutamine (Fig. S7a,b), contrasting with findings in other species where NIGT1 genes were reported to respond exclusively to nitrate (Kiba et al., 2018; Maeda et al., 2018). Interestingly, the MtNIGT1s were induced to a lower extent by the addition of both nitrate and glutamine compared to either treatment alone. Examination of MtLBD19 and MtLBD23 transcript levels revealed that they did not respond to nitrate but were significantly induced by glutamine or by a combination of glutamine and nitrate (Fig. S7c,d). These results are broadly consistent with the potential involvement of nodule produced glutamine acting through LBDs and/or NIGTs to repress NRT2.1-mediated nitrate uptake. To further investigate this possibility, the MtLBDs and MtNIGTs of interest were overexpressed in hairy roots (Fig. S8a,b,d,e). Overexpression of NIGT1.1 and NIGT1.2 suppressed NRT2.1 transcript levels under 0.5 or 5 mM nitrate (Fig. S8c), consistent with their potential importance in suppressing NRT2.1 expression during nodulation, while LBD23 downregulated NRT2.1 expression only under 5 mM nitrate (Fig. S8f). We then tested whether a short-term glutamine treatment, which repressed NRT2.1 expression, affects nitrate uptake. We found that a 6 h glutamine treatment significantly reduced the uptake of 15N-labeled nitrate in roots, but not in shoots (Fig. S9a). To test whether a longer-term treatment would have a similar effect, M. truncatula seedlings were treated weekly with 0.5 mM glutamine and 0.2 mM 15KNO3 over a 3-wk time period. The plants responded strongly in terms of growth relative to the controls grown on 0.2 mM 15KNO3, indicating that M. truncatula can efficiently uptake and utilize glutamine (Fig. S9b). Furthermore, the glutamine supplemented roots exhibited reduced 15NO3 uptake both in roots and in shoots (Fig. 1f). These findings support a potential role for nodule assimilates in repressing nitrate uptake by downregulation of MtNRT2.1. To further investigate the role of nodules in suppressing nitrate uptake in roots, we tested nitrate uptake in two M. truncatula nodulation-defective mutants, nodule inception (nin) and does not fix nitrogen 7 (dnf7). Loss of NIN completely blocks rhizobial infection and nodule formation (Liu & Bisseling, 2020), while the dnf7 mutant, which lacks Nodule Cysteine Rich Peptide 169, forms white nodules incapable of nitrogen fixation due to a defect in symbiosome development (Horváth et al., 2015). Following our earlier experiments, the mutants were grown under nitrogen-starved conditions (0.2 mM KNO3) with or without rhizobia inoculation, and the expression of MtNRT2.1 was assessed by quantitative real-time PCR at 3 wpi. Our results show that neither nin nor dnf7 mutants exhibited significant changes in MtNRT2.1 transcript levels (Fig. 2a–c). Additionally, rhizobial inoculation did not cause significant changes in the transcript levels of the LBDs or NIGTs of interest in either of the mutants (Fig. S10). In line with these findings, no differences in 15NO3 uptake were observed in the shoots and roots of nin and dnf7 seedlings after rhizobia inoculation, in contrast with the WT (Fig. 2d–f). In addition, we used a rhizobial mutant that is unable to fix nitrogen (ΔnifK) to inoculate M. truncatula under low nitrate, and no repression of MtNRT2.1 was found (Fig. S11). In summary, our results using both plant and rhizobium mutants suggest that functional nodules are required for the suppression of direct root nitrate uptake rather than nodule formation per se. In previous studies, it was shown that under low nitrate conditions nodule formation is reduced in nrt2.1 and nlp1 mutants (Misawa et al., 2022; Luo et al., 2023), which are deficient for nitrate uptake, suggesting that in WT plants, the limited N available is probably invested in forming nodules. This is consistent with our results, which show that NRT2.1 expression is unaffected at the early stages of nodule formation (Fig. S2). However, our results suggest that once productive nodules are established, direct nitrate uptake is suppressed, likely through glutamine or other assimilates. We further found that Lotus japonicus shows a similar repression of LjNRT2.1 in denodulated roots, suggesting that this may be a general feature of legumes (Fig. S12). The finding that nitrate uptake is reduced in nodulated plants is somewhat unexpected, given that on a per molecule basis, the cost of N2 fixation exceeds that of nitrate reduction to ammonia by 9 kJ (Gutschick, 1978). The prioritization of N fixation over nitrate uptake when nitrate availability is limited suggests that the ‘nitrate scavenging’ strategy, which entails root growth and release of exudates, etc., provides a lower return on investment for the plant than N fixation. Medicago truncatula (Gaertn.), ecotype R108 was used as the WT in this study. Medicago truncatula mutants nin-1 and dnf7-1 alleles were created by fast neutron mutagenesis in the Jemalong A17 background as previously described (Starker et al., 2006; Marsh et al., 2007). For nodulation, WT rhizobia Sinorhizobium meliloti 2011 or nitrogenase nifK-mutated rhizobia (fix-) S. meliloti 2011ΔnifK was used. Seeds were scarified using sandpaper, washed with sterilized deionized water, treated with bleach for 4 min, and then washed six times with distilled deionized water (ddH2O). After imbibition, the seeds were transferred to water agar plates and left in the dark for 3 d at 4°C, then the plates were inverted to allow roots to extend at 22°C overnight. Germinated seeds were transferred on Fahraeus plant (FP) medium (Fahräeus, 1957; plates (1% agar)) for 7 d in controlled environment rooms. Fahraeus plant medium composition: 0.7 mM KH2PO4, 0.5 mM MgSO4, 1 mM CaCl2, 50 μM Fe-EDTA, 0.8 mM Na2HPO4, 0.1 μg l−1 H3BO3, 0.1 μg l−1 MnSO4·H2O, 0.1 μg l−1 ZnSO4·7H2O, 0.1 μg l−1 CuSO4·5H2O, and 0.1 μg l−1 Na3MoO4. Seedlings were then transferred to trays containing 50 cells each, with a sterile substrate mixture of vermiculite and perlite (1 : 1). Each cell received 50 ml of FP medium, which did not normally contain nitrogen, supplemented with either 0.2 mM KNO3 or 0.2 mM K15NO3. Seedlings were grown for 5 d in a controlled environment chamber (22°C, 16 h light : 8 h dark photoperiod, c. 70% humidity) before inoculation. Sinorhizobium meliloti 2011 was cultured overnight at 28°C with shaking and diluted to an OD600 of 0.02 in dH2O. A 1 ml aliquot of this inoculum was applied to each seedling. After inoculation plants were watered weekly with 30 ml of FP medium containing either 0.2 mM KNO3 or 0.2 mM K15NO3. Roots of 0.2 mM KNO3 treated seedlings were harvested at 7, 14, and 21 d postinoculation (dpi) for quantitative real-time PCR (qPCR). This corresponds to 20 μg NO3 plant−1 d−1. Both root and shoots were harvested from 0.2 mM K15NO3 (10 atom %) treated seedlings at 21 dpi for quantification of 15N content. Nonrhizobial inoculated seedlings grown under similar conditions were used as a control. The Miyakojima MG-20 ecotype of L. japonicus was used as the WT plant. Seven-day-old plants were inoculated with or without Mesorhizobium loti NZP2235. After inoculation, plants were watered weekly with 30 ml of FP medium containing 0.2 mM KNO3. Roots were harvested at 21 dpi for (qPCR). Primers used were listed in Table S2. Full-length cDNA of MtNIGT1.1, MtNIGT1.2, MtLBD19, and MtLBD23 was obtained by PCR from M. truncatula R108 cDNA and cloned into the pUB-GFP overexpression construct (lab stock) to generate pUB-MtNIGT1.1, pUB-MtNIGT1.2, pUB-MtLBD19, and pUB-MtLBD23 constructs using the One Step Cloning Kit (Vazyme, Nanjing, China). The primers used for constructs are listed in Table S2. The control vector and overexpression constructs were introduced into M. truncatula R108 seedlings by A. rhizogenes Arqual-mediated hairy root transformation. Seedlings were then put on agar plates containing FP medium and kept upright in controlled environment rooms for 10 d. At this stage, untransformed roots were removed, and the seedlings were placed onto new FP plates supplemented with 0.5 mM KNO3 or 5 mM KNO3. Plants were grown for 2 wk to allow hairy roots to develop. The Green fluorescent protein (GFP) selection marker was used to identify transformed roots under a stereo-fluorescent microscope (Olympus SZX7). Roots that showed positive fluorescence were harvested for RNA extraction and quantitative real-time PCR. Total RNA was extracted from M. truncatula roots using the FastPure Universal Plant Total RNA Isolation Kit (Vazyme) following the manufacturer's protocol. RNA concentration and purity were determined using a DeNovix DS-11 Spectrophotometer. First-strand cDNA was synthesized from 1 μg of total RNA using the FastKing gDNA Dispelling RT SuperMix (Tiangen Biotech, Beijing, China) to remove any genomic DNA contamination. Quantitative real-time PCR was performed using the UltraSYBR Green Mixture qPCR kit (GenStar, Beijing, China) in a StepOnePlus™ Real-Time PCR System (Applied Biosystems, USA). The Medicago truncatula polyubiquitin 3 gene (Medtr4g088495) was used as an internal reference for the normalization of gene expression. Relative gene expression levels were calculated using the 2−ΔΔCt method. The primers used for qPCR are listed in Table S2. Nitrate uptake was assessed using the stable isotope 15NO3−. For short-term treatments, 14-d-old seedlings grown on FP medium were transferred to fresh hydroponic FP medium containing 0.2 mM K15NO3 (10% atom excess), either alone or supplemented with 0.5 mM glutamine, and incubated for 6 h. Following treatment, the seedlings were washed for 1 min in 0.1 mM CaSO4 and rinsed at least three times with ultrapure water. Roots and shoots were then separated for further analysis. For seedlings grown in the substrate, harvested roots were thoroughly washed with ultrapure water before root and shoot samples were collected for 15N analysis. All samples were dried at 65°C for 2 d, and the 15N content was measured using an Elemental Analyzer-Isotope Ratio Mass Spectrometer (Vario EL III-Isoprime, Elementar, Germany). The nitrogen stable isotope ratio was expressed as δ15N values, following international standards. Total RNA was extracted using TRIzol® Reagent according to the manufacturer's instructions and assessed for quality via Bioanalyser and NanoDrop, with only high-quality RNA used for sequencing. RNA-seq libraries were prepared and sequenced by Shanghai Majorbio Bio-pharm Biotechnology Co. Ltd, using Illumina-Stranded mRNA Prep, where mRNA was isolated by oligo(dT) beads and then fragmented by fragmentation buffer, and converted to cDNA using a SuperScript double-stranded cDNA synthesis kit (Invitrogen). Then the synthesized cDNA was subjected to end-repair, phosphorylation and ‘A’ base addition according to Illumina's library construction protocol. Libraries were size-selected, PCR-amplified, and sequenced on a NovaSeq 6000 sequencer (2 × 150 bp read length). The raw paired end reads were trimmed and quality controlled by fastp (Chen et al., 2018) and aligned to reference genome with hisat2 (Kim et al., 2015) software. Transcript assembly was done via StringTie (Pertea et al., 2015), and gene expression quantified using RSEM (Li & Dewey, 2011). Differentially expressed genes were identified using DESeq2 (Love et al., 2014) or DEGseq (Wang et al., 2009), DEGs with |log2FC| ≧ 1 and FDR ≤ 0.05 (DESeq2) or FDR ≤ 0.001 (DEGseq) were significantly DEGs. The raw data are archived with NCBI (BioSample accession SAMN46299312; BioProject ID We would to support to from the Shanghai and of and of the of and the of and to from the Shanghai of Plant We for the nrt2.1 and and the and performed the and the and and the and to this The raw data of RNA-seq is in NCBI with the accession and All available data are shown in the Fig. Nitrate uptake and MtNRT2.1 expression under high nitrate in denodulated roots. Fig. MtNRT2.1 expression in denodulated roots of Medicago truncatula plants at different time points after rhizobia inoculation. Fig. analysis of the denodulated roots and noninoculated control Fig. A phylogenetic of of Medicago Arabidopsis thaliana, and Fig. A phylogenetic of the from Medicago truncatula and Arabidopsis Fig. Nitrate induction of NRT2.1 is not by Fig. Transcript of and in to different nitrogen Fig. Overexpression of but not affects the expression of NRT2.1 in Medicago truncatula. Fig. Nitrate uptake and growth of WT seedlings grown with different nitrogen Fig. MtLBDs and MtNIGTs expression in nodulation-defective Fig. NRT2.1 expression in Medicago truncatula seedlings inoculated with a rhizobia Fig. NRT2.1 expression in roots of nodulated and Lotus japonicus Table The full list of differentially expressed Table Primers used for this study. is not for the content or of any Information by the than be to the The is not for the content or of any by the than be to the for the The remains with to in and in any