Nitrogen (N) and carbon (C) are essential elements for plant growth and crop yield. Thus, improved N and C utilisation contributes to agricultural productivity and reduces the need for fertilisation. In the present study, we find that overexpression of a single rice gene, Oryza sativa plasma membrane (PM) H+-ATPase 1 (OSA1), facilitates ammonium absorption and assimilation in roots and enhanced light-induced stomatal opening with higher photosynthesis rate in leaves. As a result, OSA1 overexpression in rice plants causes a 33% increase in grain yield and a 46% increase in N use efficiency overall. As PM H+-ATPase is highly conserved in plants, these findings indicate that the manipulation of PM H+-ATPase could cooperatively improve N and C utilisation, potentially providing a vital tool for food security and sustainable agriculture.
Nitrogen (N) and carbon (C) are indispensable elements for plant growth and are required in large quantities for crop production1. Crops largely obtain N from the soil as NH4+ and/or NO3− and C from the atmosphere as CO2. Synthetic N fertilisers are also applied in large amounts, with annual rates of >120 million tons worldwide2. Crops have a limited ability to utilise N3; thus, excess N is continuously lost from agricultural systems, which pollutes the environment4. In addition, the productivity of C3 plants such as rice and wheat is limited by inefficient CO2 fixation by RuBisCO during photosynthesis, due to low CO2 concentrations within the mesophyll cells of leaves. Plant biomass and crop production can be improved by the enhancement of intercellular CO2 concentration, which creates an effect similar to CO2 fertilisation5, but also emits excess CO2 into the atmosphere6. Thus, it is critically important to determine how best to enhance N and CO2 uptake by plants to improve crop production and environmental performance.
Plasma membrane (PM) H+-ATPase, a subfamily of P-type ATPases, generates a membrane potential and H+ gradient across the PM, energising multiple ion channels and various H+-coupled transporters for diverse physiological processes7,8. In previous studies, we demonstrated that the PM H+-ATPase mediates light-induced stomatal opening9,10. Overexpression of the PM H+-ATPase in guard cells significantly enhances stomatal opening, photosynthesis and, subsequently, growth in Arabidopsis thaliana, a model plant11. It remains unknown if this manipulation would be efficient in crops, such as rice, which is the staple food for three billion people worldwide12.
Unlike most terrestrial plants, paddy rice grows in flooded soils where ammonium (NH4+) ions constitute the dominant N source for root uptake13. To use NH4+ as an N source, rice roots require efficient uptake ability and corresponding assimilation capacity for NH4+. Conversely, high tissue accumulation of unassimilated NH4+ is usually negatively correlated with plant growth14,15. The assimilation of NH4+ in root cells requires a C skeleton as the substrate for the synthesis of amino acids through the glutamine synthetase (GS)/glutamate synthase (GOGAT) cycle. The assimilation of one molecule of NH4+ generates two molecules of H+ in the cytoplasm16. PM H+-ATPase facilitates the transport of various nutrients, such as nitrate, phosphate and potassium (K+)17,18, and maintains cytosolic H+ homeostasis by pumping H+ outside the cells19. In our previous study, NH4+ nutrition was found to induce upregulation of PM H+-ATPase activity in rice roots20. Recently, we determined that enhanced PM H+-ATPase activity in rice roots ensures rice growth at high NH4+ concentrations21. Therefore, we hypothesised that PM H+-ATPase may be involved in NH4+ metabolism in rice plants.
In this study, we examined the involvement of PM H+-ATPase in NH4+ uptake by rice roots and stomatal opening for CO2 uptake and photosynthesis in rice leaves, with the aim of developing a new strategy to improve rice yield and N use efficiency (NUE) via the overexpression of a single gene, Oryza sativa PM H+-ATPase 1 (OSA1).
PM H+-ATPase mediates NH4 + absorption
We first investigated the relationship between PM H+-ATPase and NH4+ uptake by rice roots. We treated rice roots with the fungal toxin fusicoccin (FC), a stimulator for PM H+-ATPase activity22, and found that the rate of 15NH4+ absorption increased by 17% in darkness and by 11% under illumination, compared to the corresponding controls (mock) (Fig. 1a). These results clearly indicate that PM H+-ATPase is involved in NH4+ uptake in rice roots. Under illumination, we also observed an additional increase in the 15NH4+ absorption rate (Fig. 1a) and induction of leaf stomatal opening for transpiration (Fig. 1b). These results suggest that enhanced transpiration in leaves also contributes to NH4+ uptake by roots. Therefore, we inferred that the overexpression of PM H+-ATPase in rice roots and/or stomatal guard cells would efficiently improve NH4+ absorption.
Phenotype of OSA1 overexpression and mutation rice lines
To evaluate the effects of PM H+-ATPase on NH4+ and CO2 uptake in rice, we focused on a typical PM H+-ATPase isoform, OSA1, and investigated the phenotypes of OSA1-overexpressing lines (OSA1-oxs, driven by the CaMV-35S promoter23, OSA1#1 to OSA1#3) (Fig. 2a), and osa1 knockout mutants (osa1-1 to osa1-3, TOS17 insertional mutants) (Fig. 3a and Supplementary Fig. 1). Compared to wild-type (WT) plants, OSA1 expression was 7.4–8.6-fold higher in roots and 3.5–5.3-fold higher in leaves of OSA1-oxs (Fig. 2c), without affecting the expression levels of other PM H+-ATPase isoforms (Supplementary Table 1). OSA1-ox plants exhibited ~40% higher H+-ATPase protein levels and ~30% higher PM H+-ATPase activity than did WT plants (Fig. 2d, e), whereas these values were reduced in osa1 mutants (Fig. 3c–e). We confirmed higher H+ extrusion from roots in OSA1-oxs (Supplementary Fig. 2) and proper localisation of overexpressed PM H+-ATPase in roots (Supplementary Fig. 3). When grown under hydroponic conditions, 4-week-old OSA1-ox lines exhibited enhanced plant growth, with 18–33% greater dry weight compared to the WT (Fig. 2a, b). By contrast, osa1 mutants exhibited 33–52% lower dry weight compared to WT plants (Fig. 3a, b). These results indicate that OSA1 (PM H+-ATPase) is a key factor regulating growth in rice. We observed no significant phenotype changes related to growth, relative OSA1 gene expression, PM H+-ATPase protein levels, stomatal opening (stomatal conductance) and photosynthesis rate in the empty vector-transformed rice under hydroponic conditions (Supplementary Fig. 4).
Overexpression of PM H+-ATPase enhanced NH4 + uptake
To understand the effects of PM H+-ATPase overexpression on N uptake, we compared the isotopic 15N (15NH4+) absorption rate between WT and OSA1-oxs (or osa1 mutants), and determined the absorption rate of 15NH4+ within 5 min by roots. 15NH4+ concentrations ranging from 0.5 to 8 mM were used to test 15NH4+ uptake via different NH4+ transport systems in rice roots. Interestingly, the 15NH4+ absorption rate in all OSA1-oxs was significantly higher than that in the WT, under both low (≤1 mM) and high (≥1 mM) NH4+ concentration conditions involving the high- and low-affinity transport systems, respectively24 (Fig. 2f). By contrast, all osa1 mutants exhibited lower 15NH4+ absorption rates under all NH4+ concentration conditions (Fig. 3f). We also examined the 15NH4+ absorption rate within 30 min under 2 mM 15NH4+. The rate of 15NH4+ absorption was 20–30% higher in OSA1-oxs than in the WT, but was markedly lower in osa1 mutants (Supplementary Fig. 5). In all rice lines, 15NH4+ absorption rates were significantly repressed by treatment with 0.35 µM vanadate, an inhibitor for PM H+-ATPase (Supplementary Fig. 5). These results confirmed that PM H+-ATPase modification in rice roots regulated NH4+ absorption. Consequently, under laboratory hydroponic conditions, total N accumulation was found to be 16–57% higher in OSA1-oxs (Fig. 2g) but lower in osa1 mutants (Fig. 3g) compared to the WT. In addition, the contents of other nutrients such as K, P, Ca, S, Fe, and Zn were also increased in OSA1-oxs and decreased in osa1 mutants compared to the WT (Supplementary Fig. 6a–c, f, j). Interestingly, total C accumulation was 21–47% higher in OSA1-oxs but lower in osa1 mutants compared to the WT (Figs. 2h and 3h). Because C is not taken up by plant roots, these results suggest that OSA1 modification influenced CO2 uptake and/or fixation in rice leaves.
PM H+-ATPase overexpression enhanced stomatal conductance and photosynthetic activity
Stomata are crucial for gas exchange, particularly for CO2 diffusion into the leaf12. Light, the most effective environmental signal for stomatal opening, then activates PM H+-ATPase10,11,25,26,27,28. PM H+-ATPase-induced hyperpolarisation in the PM of guard cells enables K+ uptake through inward-rectifying K+ channels. The accumulation of K+ and its counter ions in guard cells prompts guard-cell swelling and stomatal opening29. Therefore, we investigated stomatal phenotypes in OSA1-oxs. Representative closed and open stomata in a WT rice leaf are shown in Fig. 4a. In darkness, the level of stomatal closure in OSA1-oxs was similar to that in the WT, whereas under light, the ratio of open to closed stomata was significantly higher in OSA1-oxs (Fig. 4b). Conversely, in osa1 mutants, the ratio of open to closed stomata was significantly lower than in the WT under light treatment (Supplementary Fig. 7a). In all rice lines, stomatal opening was suppressed by the plant hormone abscisic acid (ABA) (Fig. 4b and Supplementary Fig. 7a), suggesting that ABA action was unaffected in guard cells of both OSA1-ox and osa1 mutant plants. Stomatal density, size, and shape in OSA1-oxs and osa1 mutants were comparable to those of the WT (Supplementary Fig. 8), suggesting that overexpression or mutation of PM H+-ATPase in rice had no effect on stomatal morphology or development; these results were similar to our observations in Arabidopsis thaliana12.
Given that stomatal aperture is a limiting factor for photosynthesis12,30, we examined the photosynthetic properties of OSA1-ox plants. Under saturated white light (WL) conditions, stomatal conductance in OSA1-oxs was almost double that in the WT (Fig. 4c and Supplementary Table 2), and photosynthetic rates in OSA1-oxs were 26–28% higher than in the WT (Fig. 4d and Supplementary Table 2), indicating that enhanced light-induced stomatal opening in OSA1-oxs conferred higher photosynthesis rates. By contrast, osa1 mutants exhibited 22–37% lower stomatal conductance and 27–35% lower photosynthetic rates (Supplementary Fig. 7b, c). Next, we examined photosynthetic light response curves in detail. Along with increased stomatal conductance (Fig. 4e), the photosynthetic rates of OSA1-ox plants were 15–34% higher than those of the WT (Fig. 4f), particularly under high-intensity light (500–1500 µmol m−2 s−1). Photosynthetic CO2 response curves (A–Ci curves) were also higher for OSA1-oxs than for the WT (Fig. 4g), indicating a higher photosynthetic capacity among OSA1-ox plants. The water use efficiency of OSA1-oxs was 13–21% lower than that of the WT (Supplementary Table 2).
Genome-wide effect of OSA1 on gene expression
To identify differentially expressed genes (DEGs) and associated pathways that may provide a molecular basis for the described OSA1-ox and osa1 mutant phenotypes, we analysed the comprehensive gene expression profiles in the leaves and roots of 4-week-old WT, OSA1-ox (OSA1#2) and osa1-2 mutant plants using RNA-sequencing (RNA-seq) analysis. Among the DEGs, 1373 and 1124 transcripts were upregulated in the leaves and roots of the OSA1-ox line, and 347 and 3295 transcripts were downregulated in the leaves and roots of the osa1-2 mutant, respectively (Fig. 5a). By contrast, 1895 and 1304 transcripts were downregulated in the leaves and roots of the OSA1-ox line, and 1859 and 2913 transcripts were upregulated in the leaves and roots of the osa1-2 mutant, respectively (Supplementary Fig. 9a–c). Consistent with OSA1 expression levels, we detected 59 and 82 genes in the leaves and roots, respectively, that were upregulated in the OSA1-ox line but downregulated in the osa1-2 mutant (Fig. 5a).
We then performed Gene Ontology (GO) term enrichment analysis of the DEGs upregulated in the OSA1-ox line and downregulated in the osa1-2 mutant to investigate the molecular mechanisms underlying OSA1-mediated biological processes (Supplementary Fig. 9d, e and Supplementary Data 1). The results indicated that 12 biological processes were significantly enriched, including photosynthesis, NH4+ assimilation, glutamate biosynthesis, amino acid metabolism, carbohydrate transmembrane transport, various ion transport, and N utilisation (Supplementary Fig. 9d, e and Supplementary Data 1). Genes associated with transmembrane transporter activity, ion transport, substrate-specific transmembrane transporter activity, cation transmembrane transporter activity, carbohydrate transmembrane transporter activity, and PM part were also significantly enriched in overlapping genes that were upregulated in OSA1-ox roots and leaves, but downregulated in the osa1-2 mutant (Supplementary Data 2). In addition, we compared leaf and root transcriptomes between the WT, OSA1-ox line, and osa1-2 mutant, and found that genes associated with nucleic acid binding transcription factor activity, response to chitin, response to organonitrogen compound, regulation of N compound metabolic process, regulation of nucleobase-containing compound metabolic process, and RNA biosynthetic process were significantly enriched in the overlapping genes downregulated in OSA1-ox leaves and upregulated in osa1 mutant leaves (false discovery rate [FDR] < 0.05) (Supplementary Data 3). However, no GO terms were found to be significantly enriched in overlapping genes downregulated in OSA1-ox roots and upregulated in osa1-2 mutant roots (Supplementary Data 3).
A set of genes were enriched in “membrane transport” category. Six NH4+ transporter genes were enriched in the membrane transport category: AMT3;3, AMT3;1, AMT2;3, AMT2;1, AMT1;2 and AMT1.1 (Fig. 5b). These genes encode both high- and low-affinity NH4+ transporters and were significantly upregulated in the OSA1-ox line and downregulated in the osa1 mutant in both leaves and roots (Fig. 5b). Transporter genes encoding other cation transporters (e.g. HAK1, CAX1a and CAT1), electroneutral substance transporters (e.g. PIP1;3) or anion transporters (e.g. PT8) were also affected by the modification of OSA1 (Fig. 5b). These results suggest a potential role for OSA1 in modulating ion and solute transport in plants.
Genes involved in NH4+ assimilation such as GS (GS2 and GS1;2) and glutamate synthase (NADH-GOGAT2, Fd-GOGAT2 and NADH-GOGAT1) were also strongly affected by the OSA1 modification (Fig. 5b). Genes associated with photosynthesis were induced by OSA1 overexpression and repressed by OSA1 knockout in leaves; these included Psb28, PsbQ, PsaH, PFPA2, PsbR1, GLO4 and RbcS (Fig. 5b).
We examined the expression levels of NH4+-responsive genes, including AMT1;1, GS1;2, NADH-GOGAT1, NADH-GOGAT2 and GS2 using quantitative reverse-transcription polymerase chain reaction (PCR) (Fig. 5d–h). The expression of all investigated genes increased significantly in the OSA1-ox lines. Notably, GRF4, a key transcription factor in N metabolism and C fixation in rice31, was highly expressed in response to OSA1 overexpression (Fig. 5c).
Overexpression of PM H+-ATPase promoted field production
To verify the effects of OSA1 overexpression on rice yield under field conditions, we conducted trials over two growing seasons at three different locations in the middle of China (Nanjing-S in 2016, and Nanjing-N and Fengyang in 2017). We applied urea as an N fertiliser at four different levels: 0 kg ha−1 (no N [N–N]), 100 kg ha−1 (low N [L–N]), 200 kg ha−1 (moderate or normal N [M–N]) and 300 kg ha−1 (high N [H–N]). Rice seedlings were planted at a spacing of 25 cm between rows and 20 cm between hills, with a total area of 26 m2 per transect. Stomatal conductance and photosynthetic rates during the vegetative stage exhibited similar trends in the field and laboratory (Supplementary Fig. 10). Representative plants at the reproductive stage under M–N conditions at Nanjing-N are shown in Fig. 6a, b. At all three locations, grain yield of the OSA1-ox lines was 27–39% (mean, 33%) higher than that of the WT (Fig. 6e and Supplementary Tables 3–5). Conversely, in osa1 mutants, grain yield was significantly lower than that of the WT at all three locations (Supplementary Tables 3–5). In OSA1-oxs, the higher yield was correlated with higher panicle weight (18–42%) (Fig. 6f), which was attributed to increased numbers of panicles per hill (15–20%) (Fig. 6c, g) and spikelets per panicle (8–16%) (Fig. 6d, h). Plant height, panicle length, filled grain rate and 1000-grain weight were nearly identical between OSA1-ox, osa1 mutant and WT plants (Supplementary Tables 3–5). Similar patterns were observed across fertilisation levels (Fig. 6j, Supplementary Fig. 11 and Supplementary Tables 3–5). Notably, under N–N conditions, grain yield was 12–20% higher in OSA1-oxs than in the WT at all test locations (Fig. 6j and Supplementary Tables 3–5). The NUE of OSA1-oxs was ~46% higher than that of the WT at all N fertilisation levels (Fig. 6i). Even when treated with only half the amount of N fertiliser (L–N, 100 kg ha−1), the grain yield of the OSA1-ox lines was significantly higher than that of the WT grown under M–N conditions (200 kg ha−1) (Fig. 6j). Thus, the same grain yield was attained using only half the amount of N fertiliser when the WT was replaced with OSA1-oxs.
To further verify the practical outcome of OSA1 overexpression in rice, we conducted an independent field trial in Hainan, southern China, which has a tropical climate and short-day conditions, and therefore produces lower yield than the subtropical areas of central China. Under these conditions, the OSA1-ox lines also produced significantly higher grain yield than the WT (Supplementary Table 6).
Increasing crop yield by improving NUE and C fixation is important for sustainable agriculture and environment performance. In this study, we demonstrated the critical role of the PM H+-ATPase gene OSA1 in controlling both NUE and photosynthesis in paddy rice production. Overexpression of OSA1 in rice plants increased the activity of PM H+-ATPase (Fig. 2e), promoted NH4+ uptake and assimilation in roots (Fig. 2f, g) and enhanced light-induced stomatal opening and stomatal conductance and photosynthetic rate under saturated WL in leaves (Fig. 4b–d and Supplementary Table 2), leading to higher NUE and grain yield (Fig. 6). Our results demonstrate the cooperative enhancement of NH4+ metabolism, photosynthesis rate and grain yield through the expression modulation of a single PM H+-ATPase gene in rice plants.
PM H+-ATPase was found to regulate NH4+ uptake in rice (Fig. 1 and Supplementary Fig. 5). Furthermore, genetic evidence based on OSA1 overexpression/knockout showed that OSA1 modulation regulated the rate of NH4+ absorption by rice roots across a wide range of rhizosphere NH4+ concentrations (Figs. 2f and 3f). RNA-seq analyses revealed the upregulation of at least six NH4+ transporter genes (AMT3;3, AMT3;1, AMT2;3, AMT2;1, AMT1;2 and AMT1.1) that encode both high- and low-affinity NH4+ transporters in OSA1-overexpressing lines (OSA1-oxs), and the downregulation of these genes in the osa1 mutant (Fig. 5). These results indicate that there is a close relationship between PM H+-ATPase and NH4+ transporters in rice root cells. These coordinated expression pattern of different genes is also the fundamental mechanisms that enable OSA1-oxs rice roots to efficiently take up NH4+ in the field soils with frequently fluctuated NH4+ concentration. Our results also indicate that OSA1 overexpression may enhance NH4+ assimilation capacity. Genes responsible for NH4+ assimilation such as glutamine synthetase (GS1;2 and GS2) and glutamate synthase (NADH-GOGAT1, NADH-GOGAT2 and Fd-GOGAT) were upregulated in OSA1-oxs (Fig. 5b, e–h). Because NH4+ uptake and assimilation are closely synchronised in plant roots32, enhanced GS and GOGAT activity can transfer root-absorbed NH4+ to amino acids for the synthesis of various N-containing compounds during plant growth and development, which in turn prevent NH4+ overloading in the root cytoplasm due to the acceleration of NH4+ uptake in OSA1-oxs (Fig. 2f). However, the process of NH4+ assimilation also generates H+, which is toxic if excessively accumulated in the cytoplasm33,34. Notably, the overexpression of only NH4+ transporter genes (AMT1;1 and AMT1;3)35,36 or glutamine synthetase (GS1;1 and GS1;2)37 alone led to higher NH4+ uptake or assimilation rates, but caused poor growth and yields in paddy rice. Considering the important role of PM H+-ATPase in maintaining intracellular pH, enhanced PM H+-ATPase activity through OSA1 overexpression (Fig. 2c–e) pumped excessive H+ out of root cells during NH4+ assimilation (Supplementary Fig. 12). Through this feedback, OSA1-ox rice absorbs and uses NH4+ more efficiently than do WT plants (Figs. 2f–g and 3f–g), which is important for NUE improvement.
We also observed that OSA1 is involved in C fixation through the regulation of stomatal opening (Supplementary Fig. 12). Stomatal conductance and photosynthetic rates were enhanced in OSA1-oxs due to increased stomatal aperture opening (Fig. 3), compared with rates in the WT and osa1 mutants (Supplementary Fig. 7). This result is consistent with the finding that genes related to photosynthesis were upregulated by OSA1 overexpression and downregulated by OSA1 knockout (Fig. 5b), for example, Psb28, PsbQ, PsaH, PFPA2, PsbR1, GLO4 and RbcS38,39,40. Enhanced photosynthesis in OSA1-oxs might also have contributed to the uptake and assimilation of NH4+ in rice roots by providing more C skeletons and energy for NH4+ metabolism processes41. Therefore, N acquisition and photosynthetic activity are intrinsically linked through overall N and C status in rice plants42, resulting in a globally coordinated increase in C and N accumulation in rice plants through OSA1 overexpression (Supplementary Fig. 13). Together, these results show that OSA1 overexpression promotes both C and N uptake and assimilation, which further regulate the expression of genes involved in C and N metabolism and contribute to plant growth and grain yield.
PM H+-ATPase is also involved in the uptake of various nutrients from plant roots by providing proton motive force17,18,43. In this study, the stronger acidification in OSA1-ox rice roots (Supplementary Fig. 2) could provide higher proton motive force in the rhizosphere for the uptake of nutrients. This is consistent with the enhanced NH4+ uptake rate in OSA1-oxs as compared with WT plants (Fig. 2f–g and Supplementary Fig. 6), and also consistent with upregulating the expression of various nutrient transporter genes, such as ammonium transporters AMT1;1/1;2/2;1/3;1/3;3, phosphate transporter PT1/PT8/PHO1.1 and potassium transporter HAK1 in roots of OSA1-oxs (Fig. 5b). These results coincided with the increased contents of N, P and K in OSA1-oxs as compared with WT plants (Fig. 2g and Supplementary Fig. 6). Recently, GRF4, a transcription factor in rice, was reported to integrate N assimilation, C fixation and plant growth; multiple N metabolism genes, such as AMT1;1, GS1;2, GS2 and NADH-GOGAT2, are positively regulated by GRF431. Here, GRF4 was found to be upregulated by OSA1 overexpression (Fig. 5b, c). It is possible that some master regulators of nutrient uptake and metabolism, such as GRF4, could be activated by OSA1 overexpression. The enhanced C fixation and N metabolism could also have a feedback on the expression of nutrient transporter genes in order to ensure sufficient supply of nutrients for the promotion of the plant growth. Further study is deserved to investigate the underlying molecular mechanisms responsible for the signal transduction initiated by OSA1 overexpression.
In contrast to CO2, which is taken up from the atmosphere, N is derived from fertilisers for most non-legume crops. Thus, cultivars with improved NUE are in urgent demand for the sustainable development of agriculture. The green revolution has boosted crop yields; however, the resulting cereal varieties are associated with reduced NUE44. Even precision crop management has led to only a slight improvement in NUE3. In this study, OSA1-ox rice exhibited both higher yield and higher NUE than the WT under a wide range of N fertilisation rates, from 0 to 300 kg N ha−1 (Fig. 6i, j and Supplementary Tables 3–5). Higher NUE in OSA1-ox rice leads to a lower demand for N fertilisers to produce similar yields of rice grain. To obtain the same yield as WT rice, OSA1-ox rice requires only half the amount of N fertiliser (Fig. 6j). This benefit will drastically reduce the cost of rice production as well as the environmental load produced by excess N accumulation due to rice production.
Given that the molecular mechanisms of nutrient uptake17,43 and light-induced stomatal opening27 are conserved in most plant species, this manipulation strategy could be applicable to many valuable crops. Therefore, we suggest designating plants overexpressing PM H+-ATPase as promotion and upregulation of plasma membrane proton-ATPase (PUMP) plants. If PM H+-ATPase overexpression can be realised using non-transgenic methods such as genome editing, these crops could have great potential for commercial use, conferring greater yields and potentially critical environmental benefits.
Seeds of WT rice (Oryza sativa L. ssp. japonica cv. Nipponbare), overexpression lines OSA1#1, OSA1#2 and OSA1#3 and mutant lines osa1-1 (TOS17 line ND3017), osa1-2 (TOS17 line ND3025), osa1-3 (TOS17 line ND3033) and CaMV-35S empty vector were surface sterilised in 10% (v:v) H2O2 for 30 min and preincubated in aerated 0.5 mM CaSO4 solution. After 2 days, all seeds were germinated on plastic support nets (mesh size, 2 mm2) floating on 1 mM CaSO4 solution for ~1 week, followed by application of IRRI (International Rice Research Institute) nutrient solution (2 mM NH4Cl, 0.5 mM K2SO4, 0.3 mM KH2PO4, 1 mM CaCl2, 1 mM MgSO4, 0.5 mM Na2SiO3·9H2O, 9 μM MnCl2, 0.39 μM Na2MoO4, 20 μM H3BO4, 0.77 μM ZnSO4, 0.32 μM CuSO4, 20 μM EDTA-Fe) at pH 5.5. For the gas-exchange experiment, seedlings of the WT, OSA1-overexpressing lines and osa1 mutants were grown in ½ IRRI nutrient solution for 1 week, followed by 5 more weeks of growth in modified IRRI nutrient solution (pH 5.5) containing 2 mM NH4Cl. The solution in the containers was replaced every 3 days. Plants for most of the laboratory experiments were grown in a greenhouse at 30 °C/24 °C (day/night) and 60–80% relative humidity. Plants for stomatal aperture and gas-exchange measurements (Fig. 4 and Supplementary Figs. 4 and 7) were grown in a growth chamber (NC-410HC, Nippon Medical & Chemical Instruments Co., Ltd, Osaka, Japan) under ~150 μmol m−2 s−1 fluorescent light at 30 °C/24 °C (12 h/12 h) and 60–80% relative humidity. The rice seeds for these experiments were of the same age.
For field experiments, plants were grown in the summer of 2016 and 2017 at four well-controlled biological experimental stations in Hainan in 2016 (N18°67′, E108°76′), southern Nanjing in 2016 (N32°01′, E118°51′), northern Nanjing in 2017 (N32°11′, E118°46′) and Fengyang in 2017 (N32°52′, E117°33′). Hainan is in a tropical monsoon zone with sandy soil, whereas the experimental sites in Nanjing and Fengyang are in a subtropical monsoon climate zone with yellow-brown soil. For each field experiment (Fig. 6, Supplementary Fig. 11 and Supplementary Tables 3–6), four levels of N (urea) fertiliser were applied: 0, 100, 200 and 300 kg N ha−1 (N–N, L–N, M–N and H–N). Seeds were germinated and seedlings were grown in a greenhouse for ~1 month at the beginning of May. The rice seedlings were hand-transplanted in a flooded field with regular hill spacing. Each fertilisation treatment was performed in one plot (6.5 m × 4 m). Rice seedlings were planted in 14 rows with 20 hills per row, for a spacing of 25 and 20 cm, respectively. Each OSA1-ox, osa1 mutant and WT line was planted in three rows (excluding border hills). At the edge of each plot, the same rice line of inside neighbour was also planted as the border hills (red box) to avoid the margin effects on the rice growth inside the plot. Each plot contained 480 hills, for a total of 1920 hills. Each field experiment consisted of four plots with different N fertilisation levels. Prior to seedling transplantation, the paddy field was fertilised with 80 kg P ha−1 as Ca(H2PO4)2 and 110 kg K ha−1 as K2SO4. The first N fertilisation was carried out at 2 days before transplantation using 33.3% of the total amount of N fertiliser, which was mixed into the soil. At the tillering stage (~1 week after transplanting), the second N fertilisation (33.3% of the total N) was carried out. The final N application (33.3% of the total N) was conducted 4 weeks later. The plant growth period (transplantation to harvest) differed among rice lines and N levels, as follows. At 0 or 100 kg N ha−1, the growth period was 109 ± 3 days for the WT and osa1 mutant and 102 ± 2 days for the overexpression lines; at 200 or 300 kg N ha−1, the growth period was 119 ± 2 days for the WT and osa1 mutant and 112 ± 2 days for the overexpression lines.
Grain yield was determined at harvest in October. At maturity, 6–8 hills of plants from each rice line were randomly selected at the centre of the plot from among a 22 × 18 array of hills (excluding the border hills) and harvested. Yield and its components were determined45,46 with minor modifications. The samples were divided into grain and straw for nutrient content analysis. Agronomic NUE was defined as the yield increase per kg N fertiliser in the field experiment. Relative agronomic NUE (Fig. 6i) was calculated as the ratio to WT rice under L–N treatment in each field trial. Statistical analyses were performed using two-tailed Student’s t tests and one-way analysis of variance followed by Tukey’s test.
Construction of the overexpression vector and transgenic plants
The open-reading frame of OSA1 was amplified using gene-specific primers (Supplementary Table 7). The fragment was treated with restriction enzymes, inserted into vectors and sequenced before transformation. Embryonic rice (O. sativa) calli were transformed via Agrobacterium-mediated transformation47. Three independent homozygous T2 or T3 lines (OSA1#1, OSA1#2 and OSA1#3) were used for all phenotypic analyses.
Quantitative reverse-transcription PCR
Total RNA was isolated from the roots of WT and transgenic plants using TRIzol reagent according to the manufacturer’s instructions48 (Invitrogen Life Technologies, Carlsbad, CA, USA). Quantitative PCR was performed using an SYBR Premix Ex Taq II (Perfect Real Time) Kit (TaKaRa Biotechnology, Dalian, China) on a Step One Plus Real-Time PCR System (Applied Biosystems, Bio-Rad, CA, USA), and the data were analysed using the 2−ΔΔCT method. The OsActin and OsGAPDH genes were used as internal references to normalise the test gene expression data. All analyses were repeated at least three times. PCR primer sets for gene amplification are listed in Supplementary Table 7.
Leaf and root samples were harvested separately. The samples were immediately homogenised in liquid N and then in ice-cold homogenisation buffer with a mortar and pestle49. The membrane proteins were collected by centrifugation and subjected to sodium dodecyl sulfate–polyacrylamide gel electrophoresis and immunoblot analysis; PM H+-ATPase was detected using anti-H+-ATPase antibody50. Actin was used as an internal control protein and was detected using anti-actin antibodies (1:3000 dilution, Sigma-Aldrich, St. Louis, MO, USA, Cat#057M4548). Relative PM H+-ATPase levels (Figs. 2d and 3d and Supplementary Fig. 4c) were estimated from the ratio of the signal intensity of PM H+-ATPase to that of actin from the same sample. WT and OSA1-ox seedlings were grown in IRRI nutrient solution containing 2 mM NH4+ for 3 weeks in a growth chamber. Immunohistochemical detection of PM H+-ATPase was performed9 (Supplementary Fig. 3). Roots of 3-week-old seedlings were harvested separately and placed in fixation buffer (4% paraformaldehyde, 60 mM sucrose and 50 mM cacodylic acid; pH 7.4) for 2 h at room temperature. Fixed samples were washed five times with phosphate-buffered saline (PBS) and embedded in 5% agar dissolved in PBS. Sections of 100-mm thickness were prepared using a vibratome (ZERO1; Dosaka EM, Kyoto, Japan) and placed on a glass slide. Samples were treated with enzyme solution (0.1% pectolyase and 0.3% Triton X-100 in PBS) for 2 h, washed five times with PBS, washed once with blocking solution (5% bovine serum albumin) for 10 min and incubated with primary antibody (anti-H+-ATPase) diluted 1000-fold in PBS overnight. On the second day, samples were washed five times with PBS, washed in blocking solution for 10 min and incubated with secondary antibody (Alexa 546, diluted 1000-fold in PBS) for 2 h. Finally, the samples were observed under confocal laser scanning microscopy (FV-10i; Olympus, Tokyo, Japan).
Measurement of PM H+-ATPase activity
We determined ATP hydrolytic activity of PM H+-ATPase51. Root and leaf tissues were ground in ice-cold homogenisation buffer to isolate the PM in a two-phase partitioning method51. PM H+-ATPase hydrolysis activity was determined as the difference between assay results with and without the addition of 0.1 mM Na3VO4 to the reaction solution (Figs. 2e and 3e). The assay was performed in 0.5 mL of reaction solution containing 30 mM BTP/MES, 5 mM MgSO4, 50 mM KCl, 50 mM KNO3, 1 mM Na2MoO4, 1 mM NaN3, 0.02% (w/v) Brij 58, and 5 mM disodium-ATP (substrate for PM H+ ATPase). The reaction was initiated by adding 30 µL of a membrane vesicle suspension containing 1–2 µg total protein and proceeded for 30 min at 30 °C; thus, inorganic phosphate was liberated after the hydrolysis of ATP. The reaction was stopped by adding 1 mL reagent (2% [v/v] concentrated H2SO4, 5% [w/v] sodium dodecyl sulfate and 0.7% [w/v] (NH4)2MoO4), followed by 50 µL 10% (w/v) ascorbic acid. After 10 min, 1.45 mL arsenite-citrate reagent (2% [w/v] sodium citrate, 2% [w/v] sodium arsenite and 2% [w/v] glacial acetic acid) was added52. Colour development was completed after 30 min and measured spectrophotometrically at 720 nm. In each test, H+-ATPase activity was calculated as the amount of phosphate liberated within 30 min mg−1 membrane protein in excess of the boiled-membrane protein control.
15N absorption rates in roots of WT, OSA1-overexpressing and osa1 plants
WT, OSA1-ox and osa1 mutant seedlings were grown in IRRI nutrient solution containing 2 mM NH4+ for 4 weeks in a growth chamber (NC-410HC, Nippon Medical & Chemical Instruments Co., Ltd.) under ~150 μmol m−2 s−1 fluorescent light at 30 °C/24 °C (12 h/12 h) and 60–80% relative humidity. To determine 15NH4+ absorption rates within 30 min, seedlings were rinsed in 0.1 mM CaSO4 for 1 min, transferred to modified IRRI nutrient solution containing 2 mM (15NH4)2SO4 (atom% 15N: 98%) incubated with mock 5 μM FC (Fig. 1a) or 350 μM vanadate (Supplementary Fig. 5) for 30 min53 and rinsed again with 0.1 mM CaSO4 for 1 min48. To determine 15NH4+ absorption rates within 5 min in roots of WT, OSA1-ox and osa1 mutant plants under different NH4+ concentrations, seedlings were incubated with 0.5, 1, 2, 4 and 8 mM 15NH4+ for 5 min (Figs. 2f and 3f).
Roots and shoots were separated for weighing, and then immediately frozen in liquid N2. After grinding, an aliquot of the powder was dried to a constant weight at 70 °C, and 10 mg of each sample was analysed using the MAT253-Flash EA1112-MS system (Thermo Fisher Scientific, Inc., USA). Each experiment was performed with three independent biological replicates, and statistical analyses were performed using two-tailed Student’s t tests.
Nutrient element analysis of plant samples
Roots and leaves were harvested from 6-week-old plants, washed three times with tap water and rinsed twice (5 min each) with deionised water to remove any adhering nutrients. The leaves and roots were dried in a forced-air oven at 70 °C for ~48 h to a constant weight for dry weight measurements (Figs. 2b and 3b and Supplementary Fig. 4a). The dried samples were ground and passed through a 1.0-mm screen. Total N/C contents (Figs. 2g, h and 3g, h) were determined via the dry combustion method using an Element Analyser (vario EL, Elementar, Langenselbold, Germany). For the analysis of mineral elements, the dry biomass was digested in H2SO4 or HClO4. P concentrations were determined using the molybdate yellow method and K concentrations were determined by flame emission photometry (Supplementary Fig. 6)54,55. The other nutrient elements were measured by ICP (Agilent 710 ICP-OES). At least three plants per treatment were harvested, and three independent biological replicates were analysed for each treatment.
Stomatal observation and quantification were performed10. Briefly, epidermal fragments were obtained by homogenising 1-week-old rice seedlings that had been grown in ½ Murashige and Skoog agar medium using a Waring blender. After passing the tissue through a 58-µm nylon mesh, the material was incubated in observation buffer containing 50 mM KCl, 0.1 mM CaCl2 and 5 mM MES-BTP with pH 6.5. After ~3-h incubation in darkness or light (150 µmol photon m−2 s−1 red light [LED-R; EYELA] plus 50 µmol photon m−2 s−1 blue light [Stick-B-32]) in 20 µM ABA, epidermal fragments were collected for microscopic observation. The percentage of open stomata (Fig. 4b and Supplementary Figs. 7a and 8c) was quantified as the number of open stomata per total stomata observed. At least 100 stomata were observed per treatment; three biological replicates were analysed for each treatment, and statistical analysis was conducted using two-tailed Student’s t tests.
Gas-exchange measurements were performed using the LI-6400 System (Li-Cor) with a standard chamber. Light and CO2 response curves were constructed based on data obtained using measurement processes and light sources12,56. The flow rate, leaf temperature and relative humidity were kept constant at 500 μmol s−1, 24 °C and 60–75% (Pa/Pa), respectively. Under each light/CO2 condition, photosynthetic rate and stomatal conductance data were collected after these values reached a steady state (15–30 min). Fully expanded leaves from 6-week-old plants were used in these experiments (Figs. 1b and 4 and Supplementary Figs. 7b, c and 4d, e). WL was provided by a fibre optic illuminator with a halogen projector lamp (15 V/150 W, Moritex, San Jose, CA, USA) as a light source powered by an MHAB-150W (Moritex) power supply. For CO2 response curves, leaves were measured at saturating WL conditions (~1500 μmol m−2 s−1) (Fig. 4g and Supplementary Fig. 7b, c). To obtain the stomatal conductance and CO2 assimilation rate data shown in Fig. 4c, d and Supplementary Table 2, leaves were measured under saturating WL conditions (~1000 μmol m−2 s−1).
For field gas-exchange measurements (Supplementary Fig. 10), the flow rate of the Li-6400 system was kept constant at 500 μmol s−1 at a leaf temperature and relative humidity of 28 °C and 40–50% (Pa/Pa), respectively. All measurements were performed before the heading stage. At least three plants were selected for measurement, and three biological replicates were analysed for each treatment.
Stomatal density and size
Three to four fully expanded leaves of 6-week-old rice plants were selected. At least five microphotographs were randomly taken of the adaxial or abaxial surface of the leaf lamina. The average stomatal density and size (long axis of each stoma) were calculated57 (Supplementary Fig. 8a, b).
High-throughput RNA-seq analysis
For RNA-seq analysis, total RNA was extracted from leaves and roots collected from 4-week-old WT, OSA1-ox (OSA1#2) and osa1 (osa1-2) mutant rice lines using a TRIzol Plus RNA Purification Kit (Thermo Fisher Scientific, Waltham, MA, USA). Complementary DNA libraries were constructed using a TruSeq RNA Sample Prep Kit v. 2 (Illumina, San Diego, CA, USA) and sequenced using a NextSeq 500 system (Illumina). Base-calling of sequence reads was performed using the NextSeq 500 pipeline software. Only high-quality sequence reads (50 continuous nucleotides with quality values >25) were used for mapping (Fig. 5a, b and Supplementary Fig. 9). Reads were mapped to O. sativa (IRGSP v. 1.0 2019.8.29) transcripts using the Bowtie software58. Experiments were repeated three times separately. We obtained 10.1–13.6 million sequence reads per experiment. Gene expression values were reported in RPM (reads per million mapped reads) units. Normalisation of read counts and statistical analyses were performed using the EdgeR software package59,60 and the Degust Ver. 3.1.0 web tool (http://degust.erc.monash.edu). Obtained RPM values were further analysed using MS Excel software. Only genes with log2 fold change ≥1 or ≤−1, and an FDR < 0.05 were considered to be significant DEGs. GO term enrichment was conducted using GO Term Enrichment tool in the Plant Transcriptional Regulatory Map (PlantRegMap) website61 (http://plantregmap.gao-lab.org/go.php). GO category (http://geneontology.org/) FDR ≤ 0.05 was regarded as significantly enriched.
Detection of rhizosphere acidification in roots
Rhizosphere acidification in WT and OSA1-oxs roots was determined51. The roots of 7-day-old plants were thoroughly washed with deionised water and spread on an agar sheet containing 0.7% (w/v) agar, 0.02% (w/v) bromocresol purple, 2 mM NH4Cl and 1 mM CaSO4 at pH 5.6. The roots were carefully pressed into the agar to avoid damage. For visualisation of rhizosphere acidification, incubation was conducted in a growth chamber in the dark for 12 h. The relative area of rhizosphere acidification (Supplementary Fig. 2b) was estimated as a ratio to the WT area (yellow area on agar sheet; Supplementary Fig. 2a). At least three plants per treatment were harvested, and three independent biological replicates were analysed for each treatment.
Quantification of H+ extrusion rate
The H+ efflux rates from the WT and OSA1-ox rice roots were measured using the scanning ion-selective electrode technique (SIET System BIO-003A, Younger USA Science and Technology Corp., Applicable Electronics Inc., Science Wares Inc., Falmouth, MA, USA) (Supplementary Fig. 2f)15,21. Briefly, seedlings were placed in 50 mL of growth solution with 2 mM NH4+ for 12 h. Then, the rice roots of 7-day-old plants were washed with deionised water and equilibrated in the measuring solution for 10 min. The equilibrated roots were then transferred to a measuring chamber, which contained 3 mL of a solution comprising 0.2 mM CaCl2, 0.1 mM KCl, 0.1 mM NaNO3 and 0.5 g L−1 MES (2-morpholinoethanesulfonic acid sodium salt) (pH 5.7). At least three plants per treatment were analysed, and three independent biological replicates were performed.
Further information on research design is available in the Nature Research Reporting Summary linked to this article.
The authors declare that the data supporting the findings of this study are available within the paper and the Supplementary information. The RNA-seq data that support the findings of this study have been deposited in the DNA Data Bank of Japan (DDBJ) with the accession number DRA011260. Source data are provided with this paper.
Marschner, P. Marschner’s Mineral Nutrition of Higher Plants (Elsevier, 2012).
Smil, V. Detonator of the population explosion. Nature 400, 415 (1999).
Omar, P., Aula, L., Oyebiyi, F. & Raun, W. R. World cereal nitrogen use efficiency trends: review and current knowledge. Agrosyst. Geosci. Environ. 2, 180045 (2019).
Stevens, C. J. Nitrogen in the environment. Science 363, 578–580 (2019).
Kimball, B. A. Crop responses to elevated CO2 and interactions with H2O, N, and temperature. Curr. Opin. Plant Biol. 31, 36–43 (2016).
Coskun, D., Britto, D. T. & Kronzucker, H. J. Nutrient constraints on terrestrial carbon fixation: the role of nitrogen. J. Plant Physiol. 203, 95–109 (2016).
Wang, Y., Shimazaki, K. & Kinoshita, T. Multiple roles of the plasma membrane H+-ATPase and its regulation. Enzymes 35, 191–211 (2014).
Falhof, J., Pedersen, J. T., Fuglsang, A. T. & Palmgren, M. Plasma membrane H+-ATPase regulation in the center of plant physiology. Mol. Plant 9, 323–337 (2016).
Toda, Y. et al. Oryza sativa H+-ATPase (OSA) is involved in the regulation of dumbbell-shaped guard cells of rice. Plant Cell Physiol. 57, 1220–1230 (2016).
Yamauchi, S. et al. Plasma membrane H+-ATPase (AHA1) plays a major role in Arabidopsis thaliana for stomatal opening in response to blue light. Plant Physiol. 171, 2731–2743 (2016).
Wang, Y. et al. Overexpression of plasma membrane H+-ATPase in guard cells promotes light-induced stomatal opening and enhances plant growth. Proc. Natl Acad. Sci. USA 111, 533–538 (2014).
Woolston, C. Rice. Nature 514, 49 (2014).
Watanabe, I. & Inubushi, K. Dynamics of available nitrogen in paddy sinks I. Changes in available N during rice cultivation and origin of N. Soil Sci. Plant Nutr. 32, 35–37 (1986).
Balkos, K. D., Britto, D. T. & Kronzucker, H. J. Optimization of ammonium acquisition and metabolism by potassium in rice (Oryza sativa L. cv. IR-72). Plant Cell Environ. 33, 23–34 (2010).
Zhang, M. et al. Involvement of plasma membrane H+-ATPase in the ammonium nutrition response of barley roots. J. Plant Nutr. Soil Sci. 181, 878–885 (2018).
Maschaux-Daubresse, C. et al. Nitrogen uptake, assimilation and remobilization in plants: challenges for sustainable and productive agriculture. Ann. Bot. 105, 1141–1157 (2010).
Wang, E. et al. A H+-ATPase that energizes nutrient uptake during mycorrhizal symbioses in rice and Medicago truncatula. Plant Cell 26, 1818–1830 (2014).
Briskin, D. & Gawienowski, M. Role of the plasma membrane H+-ATPase in K+ transport. Plant Physiol. 111, 1199–1207 (1996).
Falhof, J., Pedersen, J. T., Fuglsang, A. T. & Palmgren, M. G. Plasma membrane H+-ATPase regulation in the center of plant physiology. Mol. Plant 9, 323–337 (2016).
Zhu, Y. et al. Adaptation of plasma membrane H+-ATPase of rice roots to low pH as related to ammonium nutrition. Plant Cell Environ. 32, 1428–1440 (2009).
Weng, L. et al. Potassium alleviates ammonium toxicity in rice by reducing its uptake through activation of plasma membrane H+-ATPase to enhance proton extrusion. Plant Physiol. Biochem. 151, 429–437 (2020).
Kinoshita, T. & Shimazaki, K. Analysis of the phosphorylation level in guard-cell plasma membrane H+-ATPase in response to fusicoccin. Plant Cell Physiol. 42, 424–432 (2001).
Odell, J. T., Nagy, F. N. & Chua, H. Identification of DNA sequences required for activity of the cauliflower Mosaic Virus35S promoter. Nature 313, 810–812 (1985).
Sonoda, Y. et al. Distinct expression and function of three ammonium transporter genes (OsAMT1;1–1;3) in rice. Plant Cell Physiol. 44, 726–734 (2003).
Shimazaki, K., Doi, M., Assmann, S. M. & Kinoshita, T. Light regulation of stomatal movement. Annu. Rev. Plant Biol. 58, 219–247 (2007).
Kinoshita, T. & Shimazaki, K. Blue light activates plasma membrane H+-ATPase by phosphorylation of the C-terminus in stomatal guard cells. EMBO J. 18, 5548–5558 (1999).
Kinoshita, T. et al. phot1 and phot2 mediate blue light regulation of stomatal opening. Nature 414, 656–660 (2001).
Ando, E. & Kinoshita, T. Red light-induced phosphorylation of plasma membrane H+-ATPase in stomatal guard cells. Plant Physiol. 178, 838–849 (2018).
Inoue, S. & Kinoshita, T. Blue light regulation of stomatal opening and the plasma membrane H+-ATPase. Plant Physiol. 174, 531–538 (2017).
Farquhar, G. D. & Sharkey, T. D. Stomatal conductance and photosynthesis. Annu. Rev. Plant Physiol. 33, 317–345 (2003).
Li, S. et al. Modulating plant growth–metabolism coordination for sustainable agriculture. Nature 111, 595–560 (2018).
Xu, G. H., Fan, X. R. & Miller, A. J. Plant nitrogen assimilation and use efficiency. Annu. Rev. Plant Biol. 63, 153–182 (2012).
Britto, D. T. & Kronzucker, H. J. Nitrogen acquisition, PEP carboxylase, and cellular pH homeostasis: new views on old paradigms. Plant Cell Environ. 28, 1396–1409 (2005).
Feng. H., Fan. X., Miller. A. J. & Xu. G. Plant nitrogen uptake and assimilation: regulation of cellular pH homeostasis. J. Exp. Bot. https://doi.org/10.1093/jxb/eraa150 (2020).
Hoque, M. H. et al. Over-expression of the rice OsAMT1-1 gene increases ammonium uptake and content, but impairs growth and development of plants under high ammonium nutrition. Funct. Plant Biol. 33, 153–163 (2006).
Li, C. et al. The OsAMT1.1 gene functions in ammonium uptake and ammonium–potassium homeostasis over low and high ammonium concentration ranges. J. Gen. Genom. 43, 639–949 (2016).
Cai, H. et al. Overexpressed glutamine synthetase gene modifies nitrogen metabolism and abiotic stress responses in rice. Plant Cell Rep. 28, 527–537 (2009).
Xin, W. et al. An integrated analysis of the rice transcriptome and metabolome reveals differential regulation of carbon and nitrogen metabolism in response to nitrogen availability. Int. J. Mol. Sci. 20, 2349 (2019).
Lu, Y. et al. Suppression of glycolate oxidase causes glyoxylate accumulation that inhibits photosynthesis through deactivating Rubisco in rice. Physiol. Plant. 150, 463–476 (2014).
Suzuki, Y., Makino, A. & Mae, T. Changes in the turnover of Rubisco and levels of mRNAs of rbcL and rbcS in rice leaves from emergence to senescence. Plant Cell Environ. 24, 1353–1360 (2001).
Ariz, I. et al. Changes in the C/N balance caused by increasing external ammonium concentrations are driven by carbon and energy availabilities during ammonium nutrition in pea plants: the key roles of asparagine synthetase and anaplerotic enzymes. Physiol. Plant. 148, 522–537 (2013).
Coruzzi, G. M. & Zhou, L. Carbon and nitrogen sensing and signaling in plants: emerging “matrix effects”. Curr. Opin. Plant Biol. 4, 247–253 (2001).
Palmgren, M. G. Plant plasma membrane H+-ATPase: powerhouses for nutrient uptake. Annu. Rev. Plant Physiol. Plant Mol. Biol. 52, 817–854 (2001).
Gooding, M. J., Addisu, M., Uppal, R. K., Snape, J. W. & Jones, H. E. Effect of wheat dwarfing genes on nitrogen-use efficiency. J. Agric. Sci. 150, 3–22 (2012).
Chen, J. et al. pOsNAR2.1:OsNAR2.1 expression enhances nitrogen uptake efficiency and grain yield in transgenic rice plants. Plant Biot. J. 15, 1273–1283 (2017).
Fan, X. et al. Overexpression of a pH-sensitive nitrate transporter in rice increase crop yields. Proc. Natl Acad. Sci. USA 113, 7118–7123 (2016).
Hiei, Y., Ohta, S., Komari, T. & Kumashiro, T. Efficient transformation of rice (Oryza sativa L.) mediated by Agrobacterium and sequence analysis of the boundaries of T-DNA. Plant J. 6, 271–282 (1994).
Tang, Z. et al. Knockdown of a rice stelar nitrate transporter alters long distance translocation but not root influx. Plant Physiol. 160, 2052–2063 (2012).
Hayashi, M., Inoue, S., Takahashi, K. & Kinoshita, T. Immunohistochemical detection of blue light-induced phosphorylation of plasma membrane H+-ATPase in stomatal guard cells. Plant Cell Physiol. 52, 1238–1248 (2011).
Hayashi, Y. et al. Biochemical characterization of in vitro phosphorylation and dephosphorylation of the plasma membrane H+-ATPase. Plant Cell Physiol. 51, 1186–1196 (2010).
Yan, F., Zhu, Y., Müller, C., Zörb, C. & Schubert, S. Adaptation of H+-pumping and plasma membrane H+-ATPase activity in proteoid roots of white lupin under phosphate deficiency. Plant Physiol. 129, 50–63 (2002).
Baginski, E., Foa, P. & Zak, B. Determination of phosphate: study of labile organic phosphate interference. Clin. Chim. Acta 15, 155–158 (1967).
Zhu, Y., Yan, F., Zörb, C. & Schubert, S. A link between citrate and proton release by proteoid roots of white lupin (Lupinus albus L.) grown under phosphorus-deficient conditions? Plant Cell Physiol. 46, 892–901 (2005).
Chang, C. et al. Proton pump OsA8 is linked to phosphorus uptake and translocation in rice. J. Exp. Bot. 60, 557–565 (2009).
Shen, Z. et al. Induced soil microbial suppression of banana fusarium wilt disease using compost and biofertilizers to improve yield and quality. Eur. J. Soil Biol. 57, 1–8 (2013).
Wang, Y. & Kinoshita, T. Measurement of stomatal conductance in rice. Bio-protocol 7, e2226 (2017).
Wang, Y., Noguchi, K. & Terashima, I. Photosynthesis-dependent and -independent responses of stomata to blue, red and green monochromatic light: differences between the normally oriented and inverted leaves of sunflower. Plant Cell Physiol. 52, 479–489 (2011).
Langmead, B., Trapnell, C., Pop, M. & Salzberg, S. L. Ultrafast and memory-efficient alignment of short DNA sequences to the human genome. Genome Biol. 10, R25 (2009).
Robinson, M. D., McCarthy, D. J. & Smyth, G. K. EdgeR: a bioconductor package for differential expression analysis of digital gene expression data. Bioinformatics 26, 139–140 (2010).
Anders, S. et al. Count-based differential expression analysis of RNA sequencing data using R and Bioconductor. Nat. Protoc. 8, 1765–1786 (2013).
Tian, F., Yang, D., Meng, Y. Q., Jin, J. & Gao, G. PlantRegMap: charting functional regulatory maps in plants. Nucleic Acids Res. 48, D1104–D1113 (2020).
This work was supported by grants from the National Key Basic Research and Development Program (2017YFD0200200/0200206 to Y.Z.), the Natural Science Foundation of China (NSFC 31471937 to Y.Z.), Technology of Japan and the Advanced Low Carbon Technology Research and Development Program from the Japan Science and Technology Agency (JPMJAL1011 to T.K.) and the Natural Science Foundation of Anhui Province, China (1608085MC59 to X.X.), as well as Grants-in-Aid for Scientific Research on Innovative Areas (15H05956, 20H05687 and 20H05910 to T.K.).
The authors declare no competing interests.
Peer review information Nature Communications thanks Xiangdong Fu, Brent Kaiser, Beom-Gi Kin, and the other, anonymous, reviewer(s) for their contribution to the peer review of this work. Peer review reports are available.
Publisher’s note Springer Nature remains neutral with regard to jurisdictional claims in published maps and institutional affiliations.
About this article
Cite this article
Zhang, M., Wang, Y., Chen, X. et al. Plasma membrane H+-ATPase overexpression increases rice yield via simultaneous enhancement of nutrient uptake and photosynthesis. Nat Commun 12, 735 (2021). https://doi.org/10.1038/s41467-021-20964-4