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Nitrogen Journey in Plants: From Uptake to Metabolism, Stress Response, and Microbe Interaction

Omar Zayed, Omar A. Hewedy, Ali Abdelmoteleb and 5 more

Biomolecules · 2023 · Review · CC BY

doi:10.3390/biom13101443

Abstract

Plants uptake and assimilate nitrogen from the soil in the form of nitrate, ammonium ions, and available amino acids from organic sources. Plant nitrate and ammonium transporters are responsible for nitrate and ammonium translocation from the soil into the roots. The unique structure of these transporters determines the specificity of each transporter, and structural analyses reveal the mechanisms by which these transporters function. Following absorption, the nitrogen metabolism pathway incorporates the nitrogen into organic compounds via glutamine synthetase and glutamate synthase that convert ammonium ions into glutamine and glutamate. Different isoforms of glutamine synthetase and glutamate synthase exist, enabling plants to fine-tune nitrogen metabolism based on environmental cues. Under stressful conditions, nitric oxide has been found to enhance plant survival under drought stress. Furthermore, the interaction between salinity stress and nitrogen availability in plants has been studied, with nitric oxide identified as a potential mediator of responses to salt stress. Conversely, excessive use of nitrate fertilizers can lead to health and environmental issues. Therefore, alternative strategies, such as establishing nitrogen fixation in plants through diazotrophic microbiota, have been explored to reduce reliance on synthetic fertilizers. Ultimately, genomics can identify new genes related to nitrogen fixation, which could be harnessed to improve plant productivity.

Full text

From PubMed Central, U.S. National Library of Medicine, as current on Oct 23, 2026. Paragraph numbers are ours, so an answer can point at one.

1. Introduction

Soil erosion, compaction, acidification, and contamination are significant factors causing soil degradation, affecting 50% of agricultural land and food production. Consequently, 75 billion tons of fertile soil are lost worldwide due to degradation annually. Moreover, massive amounts of synthetic agro inputs to fertilize crops used by farmers harm environmental health. In this sense, applying nitrogen (N) fertilizers has increased sharply by 7.4 times compared with crop productivity, which has only increased by 2.4 times, indicating that crops have been reduced in their ability to use N efficiently, increasing food insecurity. Therefore, innovative and sustainable strategies in the agro-biotechnological sector have received more attention from botanical scientists worldwide over the last century to solve food security challenges and develop environmentally friendly products. Throughout time, microbial diversity and biomass have been assessed as partners in the plant–microbe association in maintaining soil multifunctionality.

N is a vital macronutrient for plants and a crucial component of amino acids that serve as the building blocks of enzymes and proteins in plants. Additionally, N is a part of the chlorophyll molecule, an essential factor in photosynthesis for absorbing sunlight energy, promoting plant growth and grain yield. Even the roots contain this essential element because proteins and enzymes control water and nutrient intake. Recently, Zhang et al. (2022) found that N can significantly impact flowering time. In addition to being a crucial nutrient for plant growth and development, N has a strong correlation with several abiotic stress responses. Moreover, N has a leading role in plant adaptation to the deficiency of macro- and micronutrients. For macronutrients, the processes of N and phosphorus (P) uptake interact together, creating a nutritional balance under fluctuating availability of nutrients, while potassium (K) and nitrate (NO3−) translocation and intake are positively connected as well. The plant nitrate transporters (NRTs) regulate the K translocation. Regarding micronutrients, ammonium (NH4+) application promotes iron (Fe) uptake; in contrast, NO3− triggers the chlorosis symptoms of Fe deficiency. Sulfur (S), a crucial component of enzymes involved in N metabolism, is correlated with N uptake. Alternatively, a reverse relationship between molybdenum (M) and N has been noticed. In addition, through the mechanism of anion-cation balance, approximately 90% of stress responses are influenced by NO3− uptake, which promotes the synergetic transfer of cations, including K+, sodium (Na+), cadmium (Cd2+), and zinc (Zn2+) while inhibiting the absorption of anions like chloride (Cl−) and sulfate (SO4−). Under drought stress, NO3− and NH4+ concentrations have unique impacts on plant performance. While the usage of NH4+ reduces the effects of drought on plant development, NO3− application has a reverse influence.

In this review, we aim to provide a detailed story of the N journey starting from its uptake from soil either in NO3− or NH4+ form, followed by a structural investigation of the transporters of these two forms. Upon reaching their destination, NO3− and NH4+ are exposed to various metabolic pathways (e.g., oxidation and denitrification). Next, N assimilation occurs under the intersection with glutamine synthetase (GS) and glutamate synthase or glutamine 2-oxoglutarate aminotransferase (GOGAT). Then, a comprehensive survey of the role of N in plant adaptation/tolerance to abiotic stresses, either drought or stress, is discussed. Finally, the detrimental effects of nitrogen mineral fertilization and the possibility of microbial application and genomic approaches are summarized.

2. Nitrogen Uptake

N is an essential macronutrient for plants, and it can be acquired from the soil in inorganic forms (NO3− and NH4+), which are transported across the root plasma membrane by different families of transporters. Under normal soil conditions, N is mainly available in the form of NO3−, and four families of transporters mediate NO3− uptake, namely NRT1, NRT2, chloride channel (CLC-1), and slow anion channel-associated 1 homolog 3 (SLAC1/SLAH). These transporters have distinct characteristics in affinity, capacity, regulation, and localization. NH4+ uptake is facilitated by ammonium transporters (AMTs), high-affinity transporters expressed mainly in the root hairs and epidermis. NH4+ is the dominant form of N in flooded or acidic soils, and AMTs-mediated acquisition is crucial for the N demand of plants growing in such conditions. Besides inorganic N, plants can also absorb organic N in the form of amino acids (AAs), which are abundant in soils that receive organic amendments, such as manure or compost. Several AAs transporters have been identified in plant roots, including amino acid permease (AAP1 and AAP5), proline transporter (ProT2), and lysine and histidine transporters (LHT1 and LHT6). These transporters have different substrate specificities and expression patterns contributing to the uptake of a range of amino acids from the soil solution

3.1. Plant Nitrate Transporters

Plants acquire NO3− from the soil through various transporters that belong to different families. The NRT1 and NRT2 families are the primary transporters involved in NO3− uptake, and they have distinctive roles in plant growth and seed development. The NRT1 family is a large and diverse group of nitrate peptide transporter family (NPF). There are 53 NRT1 genes in Arabidopsis and 93 in rice. These genes can be classified into 8–10 subfamilies; NRT1.1 and NRT1.2 are responsible for NO3− root uptake. The NRT1 family can transport NO3− and other substrates such as hormones, nitrite, amino acids, peptides, chloride, glucosinolates, and jasmonate-isoleucine. The NRT2 family is a smaller, more specific group of transporters expressed mainly under low NO3− conditions. In Arabidopsis, four NRT2 transporters (NRT2.1, NRT2.2, NRT2.4, and NRT2.5) function in nitrate influx and account for 95% of nitrate uptake under low NO3− concentrations. However, NRT2.1 and NRT2.2 are significant members of the NRT2 family for nitrate uptake. The NRT2 family has different spatial and temporal expression patterns, and they transport NO3− from different sources to different tissues. For example, NRT2.4 and NRT2.5 absorb NO3− from the soil root hairs, while NRT2.1 and NRT2.2 transport it from the apoplast to the cortex and endodermis cells. Moreover, NRT2.5 is induced by long-term starvation and enhances NO3− uptake from both shoots and roots of mature plants. Furthermore, another transporter family called NRT3 is also involved in NO3− transport in plants. The NRT3 family consists of two members: NRT3.1 and NRT3.2. These transporters form a complex with NRT2 transporters and regulate their activity and stability.

NO3− transceptor (transporter/receptor) NRT1.1 regulates the expression levels of many NO3− assimilation pathway genes by sensing the external NO3− concentration and modulating the root growth accordingly. NO3− transporters are regulated through phosphorylation, mediated by calcium-dependent calcineurin B-like (CBL) and CBL-interacting protein kinase (CIPK). The activity and specificity of NRT1.1 are regulated by two CIPKs, CIPK8 and CIPK23, which interact with NRT1.1 and mediate low- and high-affinity responses, respectively. At high NO3− concentrations, NRT1.1 activates the expression of Arabidopsis nitrate regulated 1 (ANR1), a transcription factor that promotes lateral root growth and proliferation. At low NO3− concentrations, NRT1.1 inhibits lateral root development by controlling auxin levels and meristem activation. To transmit the NO3− signals from NRT1.1 to the nucleus, calcium acts as a secondary messenger that modulates the expression of NO3−-responsive genes. Three calcium-dependent protein kinases (CPK10, CPK30, and CPK32) and their partner CBLs are involved in this process.

Several NRTs are involved in the loading and unloading of NO3− from the xylem and phloem, which affects the distribution and availability of NO3– within the plant. For example, NRT1.5, NRT1.8, and NRT1.9 are expressed in the xylem and phloem, and they function in the influx/efflux, removal, and loading of NO3− from/to the root vascular tissues, respectively. Other studies have shown that overexpressing NRTs in different plant species can improve N-use efficiency (NUE) as well as plant biomass to nitrogen input. However, the effects of NRT overexpression may depend on the tissue specificity, NO3– concentration, and interaction with other genes. For instance, only NRT1 transporters have been successfully overexpressed in both leaves and roots to enhance NUE, while the role of NRT2 transporters in this process is still unclear. Recently, a chimeric NRT1 transporter, AtNC4N, was overexpressed in the phloem of old leaves in Arabidopsis, rice, and tobacco, and it increased NO3– uptake and NUE under low nitrate conditions. Another NRT1 transporter, OsNRT1.1A (OsNPF6.3), was found to improve NUE, flowering time, yield, and maturity in rice. Several other NRTs, such as OsNRT1.1B (OsNPF6.3B), OsNRT2.1, OsNRT2.3a/b, OsPTR9, OsAMT1.1, and qNGR9, have also been reported to enhance NUE under high or low nitrate levels in rice.

Moreover, some genes that regulate or interact with NRTs have been identified as potential targets for improving NUE in plants. For example, the transcription factor OsNAC42 activates the expression of several NRTs and increases nitrate assimilation and NUE in rice. The nitrate reductase gene OsNR2 converts nitrate to ammonium, affecting the expression of NRTs and other nitrogen-related genes in rice. The nitrate transporter OsNPF4.5 is regulated by a microRNA (miR827) and modulates nitrate distribution and remobilization in rice. These studies demonstrate the complexity and diversity of nitrate transport and signaling in plants as well as provide valuable insights for improving NUE and crop production. Notably, NUE can be improved by transcription factors (TFs) in multiple crops. TFs are primarily proteins (rarely RNA) that control the transcription of genes by interacting with their unique promotor sequences. Numerous TF families have been found and proven to interact with particular DNA sequences on regulatory regions of NO3−-responsive genes. According to reports, these TF families act as essential regulators of the plant N response, which encompasses MADS-box, NLP, b-ZIP, LBD, NAC, MYB, GARP, TCP, AP2/ERF, and zinc/finger proteins.

NRT1, NRT2, and NRT3 have different structural conformations (Figure 1). As a result, the Pfam database collects these genes under distinct families: PF00854 for NRT1, PF07690 for NRT2, and PF16974 for NRT3. PF00854 refers to the proton-dependent oligopeptide transporter (POT) family (also known as the peptide transporter (PTR) family) and is a class of energy-dependent transporters discovered in organisms ranging from bacteria to humans. However, some family members are NO3– permeases, while others are in histidine transport. The NRT1 transporter adopts a standard major facilitator superfamily (MFS) fold, which is described by 12 transmembrane helices (TMHs) with a pseudo 2-fold axis connecting the N-terminal (TMH1–6) and C-terminal (TMH7–12) domains.

NRT2 members belong to the PF07690 (MSF) family, representing the largest family of secondary transporters, with members ranging from Archaea to Homo sapiens. MFS proteins target a diverse range of substrates in both directions across the membrane, including ions, carbohydrates, lipids, amino acids and peptides, nucleosides, and other small molecules, in many cases catalyzing active transport by converting the energy stored in a proton electrochemical gradient into a concentration gradient of the substrate. NRT3 candidates lie under the PF16974 family, which contains a C-terminal transmembrane domain required for high-affinity nitrate absorption. NRT3 is implicated in the inhibition of lateral root initiation in the presence of high sucrose-to-nitrogen ratios in the medium. As a result, the NRT2 and NRT3 component systems are believed to be implicated in the signaling pathway that integrates nutritional inputs for the regulation of lateral root architecture. In Arabidopsis, the functional unit of the high-affinity nitrate influx complex is most likely a tetramer composed of two NRT2 and NRT3 subunits each.

3.2. Plant Ammonium Transporters

As mentioned earlier, NO3− and NH4+ are the principal inorganic nitrogen sources absorbed by plant roots. Because NH4+ requires less energy than NO3− assimilation, NH4+ is the preferred form of N uptake when plants grow under N deficiency. Although NH4+ uptake is more efficient, high absorbed amounts of NH4+ by the plant can be hazardous; thus, the NH4+ uptake system in plants receives special attention. Recent research indicates that AMTs also play a role in a variety of other physiological processes such as transporting NH4+ from symbiotic fungi to plants, delivering NH4+ from roots to shoots, transmitting NH4+ in leaves and reproductive organs, and encouraging resistance to plant diseases through NH4+ transport. Aside from being a transporter, many AMTs are essential for root growth in the presence of NH4+. To avoid the adverse effects of insufficient or excessive NH4+ intake on plant growth and development, AMTs activities are fine-tuned not only at the transcriptional level through the involvement of at least four transcription factors but also at the protein level through phosphorylation, pH, and endocytosis.

The NH4+ transporter domain (PF00909) comprises two structural copies of five helices plus one extra C-terminal helix. It has been defined as an 11-times-spanning membrane channel. Ideal NH4+ transporter domain is composed of 11–12 transmembrane spots, with feature sequences “D (F Y W S) A G (G S C) X2 (L I V) (E H) X2 (G A S) (G A) X2 (G A S) (L F)” at transmembrane spot 5 and “D D X (L I V M F C) (E D G A) (L I V AC) X3 H (G A L I V) X2 (G S) X (L I V A W) G” at transmembrane region 10. AMTs can be divided into two subtypes: AMT1 and AMT2 or methylammonium permeases (MEP). Figure 1 illustrates the diversity and complexity of the N transporters in plants and their distinctive features.

3.3. Plant Nitrite Transporters

Cytosol is the first cellular compartment to encounter NO3− in the root that needs to be converted to NO2− to make organic compounds such as amino acids. The cytosolic NO2− is toxic to the cytosol and needs to be transported to the chloroplasts, where it is reduced to NH3 by NIR. Cytosolic NO2− needs to be actively transported across the membrane of the chloroplasts by a specific transporter to avoid accumulation and toxicity. Several types of plastids and plasma membrane-localized NIR have been described in Escherichia coli, Cyanobacteria, green algae, red algae, Aspergillus nidulans, diatoms, and a haptophyte, but there was no evidence of a nitrite-specific plastid and plasma membrane transporter in higher plants. Consequently, it has been hypothesized that NO3− and NO2− enter the plasma membrane via similar transporters due to previously observed competitive reduction of NO2− absorption by NO3− and vice versa.

In 1988, Brunswick and Cresswell found that illuminated chloroplasts take up NO2− in vitro. In 2007, Saguira et al. revealed that CsNitr1-L is the NO2− transporter in the chloroplasts of plants and belongs to the proton-dependent oligopeptide transporter (POT) family. CsNitr1-L, a homologue of NRT1 transporter, can transport NO2− depending on the energy availability. In 2013, Kotur et al. used 13NO2− to characterize nitrite influx and nitrite-specific transporter into roots of Arabidopsis thaliana. Fortunately, they successfully confirmed that Arabidopsis roots include a transporter specific for nitrite. Furthermore, in 2014, Maeda et al. succeeded in characterizing two genes, AT5G62720 (AtNITR2;1) and AT3G47980 (AtNITR2;2), which encode an integral membrane protein related to the NO2− transport activity from Arabidopsis, and these two genes were detected based on homologs genes from cyanobacterial genomes.

AtNITR2;1 and AtNITR2;2 are HPP-domain-containing proteins (Figure 1). These two transporters belong to PF04982, constituting integral membrane proteins with four transmembrane-spanning helices. Interestingly, their work has great value, as the HPP proteins had no functional annotation before their work. Moreover, the kinetic analyses showed that the proteins that encode from these previous two genes of Arabidopsis have a much higher affinity for NO2− than the cyanobacterial proteins. AtNITR2;1, isolated from the chloroplasts mutant lines, showed much lower NO2− uptake activity than the chloroplasts isolated from the wild-type Col-0 plants. AtNITR2;2, expressed in roots and not detectable in shoots, had a putative transit peptide similar to that of AtNITR2;1 and demonstrated low but significant activity of NO2− transport in the cyanobacterial cell. Finally, it was discovered that NO2− increased the expression of AtNITR2;1 and AtNITR2;2 under the direction of NIN-like proteins, indicating that these two proteins are nitrate-inducible parts of the nitrite transport system of plastids.

4. Nitrogen Metabolism

N is among the most widely distributed elements in the lithosphere, atmosphere, hydrosphere, and biosphere. The lithosphere contains 94% of all nitrogen on Earth (e.g., NO3−, NO2−, and NH4+), with the remaining 6% in the atmosphere (e.g., NO, N2O, and N2) and a trace (0.006%) in the hydrosphere and biosphere (e.g., NO3−, NO2−, and NH4+). N is the fourth most prevalent element in the biosphere after oxygen, carbon, and hydrogen and is an essential component of total biomass. The relative quantity of N in the biosphere reflects the relevance of N to living organisms. Although organic N, such as amino acids, hormones, and nucleic acids, is vital to life on Earth, The global N cycle is dominated by inorganic N molecules (i.e., NO3−, NO2, nitrous oxide (N2O), NO, dinitrogen (N2), and NH4+). Figure 2 summarizes all N metabolism pathways.

NO3− is the most oxidized form of N, while NH4+ is the most reduced form. Many microorganisms, mainly microbes, drive the global N cycle by metabolizing N through a variety of redox processes for energy transduction, detoxification, or assimilation via various sub-pathways such as N fixation (KEGG;M00175), assimilatory NO3− reduction (KEGG;M00531), dissimilatory NO3− reduction (KEGG;M00530), denitrification (KEGG;M00529), nitrification (KEGG;M00528), complete nitrification, complete ammonia oxidation (comammox) (KEGG;M00804), and anammox (KEGG;M00973).

Moreover, each sub-pathway from these previous sub-pathways consists of a list of genes and their encoded enzymes that are involved in the various biogenic processes in the nitrogen cycle. The nitrogen fixation path consists of nitrogenase molybdenum-iron protein alpha chain (NIFD), vanadium-dependent nitrogenase alpha chain (VNFD), and nitrogenase delta subunit (ANFG) genes.

In this sub-pathway, these previous genes were used to convert N2 to NH3 by oxidoreductase reaction. NH3 is widely used in different vital pathways for living organisms, such as arginine biosynthesis (KEGG;map00220), purine metabolism (KEGG;map00230), alanine, aspartate, and glutamate metabolism (KEGG;map00250). Also, NH3 is involved in glycine, serine, and threonine metabolism (KEGG;map00260) as well as cyanoamino acid metabolism (KEGG;map00460), glyoxylate and dicarboxylate metabolism (KEGG;map00630), and lipoic acid metabolism (KEGG;map00785). NH3 is a component in biosynthesis of secondary metabolites (KEGG;map01110), microbial metabolism in diverse environments (KEGG;map01120), carbon metabolism (KEGG;map01200), biosynthesis of amino acids (KEGG;map01230), proximal tubule bicarbonate reclamation (KEGG;map04964), collecting duct acid secretion (KEGG;map04966), protein digestion and absorption (KEGG;map04974), Vibrio cholerae infection (KEGG;map05110), and epithelial cell signaling in Helicobacter pylori infection (KEGG;map05120).

In addition, the anammox path consists of hydrazine dehydrogenase (HDH), hydrazine synthase subunit (K20932), and nitrite reductase (NO forming) (NIRK) genes, and these previous genes can convert NO2 to N2 or NH3 using nitrite reductase enzyme. In that context, the denitrification path consists of four genes: nitrous-oxide reductase (NOSZ), NIRK, nitrate reductase (cytochrome) (NAPA), and nitric oxide reductase subunit B (NORB). In this sub-pathway, the denitrification enzymes convert NO3− to N2 through NO2, NO, and N2O. Furthermore, assimilatory and dissimilatory NO3− reduction paths have different genes, such as nitrate reductase/nitrite oxidoreductase, alpha subunit (NARG, NARZ, and NXRA), NAPA, nitrite reductase (NADH) large subunit (NIRB), nitrite reductase (cytochrome C-552) (NRFA), ferredoxin-nitrate reductase (NARB), nitrate reductase (NAD-(P)-H) (NR), assimilatory nitrate reductase catalytic subunit (NASC and NASA), nitrite reductase (NAD-(P)-H) (NIT-6), ferredoxin-nitrite reductase (NIRA), and nitrite reductase (NAD(P)H) large subunit (NASD and NASB) that can convert NO2 to NH3. Figure 2 and Table 1 show the enzymes and genes involved in the N cycle.

4.1. GS and GOGAT

N uptake for plant growth has been extensively studied and discussed previously. N uptake involves an active transport across the plasma membrane of both root epidermal and cortical cells. The gene expression patterns, post-translational regulation, and the localization of NRTs and AMTs enable directed transport from the epidermis to vascular tissue and acclimation to various concentrations of N in the soil. Mostly, the plant uptakes N from the soil in the form of NO3− that is reduced by NR to NO2−, which is then imported into the chloroplast and reduced further by NIR into NH4+. NO2− reduction to NH4+ occurs in the stroma of chloroplasts and is catalyzed by the enzyme NIR. NIR reduces NO3− to NO2− using electrons from NAD(P)H, and the reduced ferredoxin (Fdred) generated due to the photosynthetic electron transport is the electron donor for NO2− reduction. NH4+ is combined with glutamate to be assimilated into glutamine. NR is regulated by NO3−, N, light, growth conditions, hormones, N metabolites, and phosphorylation.

In various biological systems, the interconversion of glutamate and glutamine plays a central role in N assimilation, transport, and recycling. GS and GOGAT are vital enzymes orchestrating these interconversions, ensuring optimal N utilization and maintaining N homeostasis. The roots are the principal site of NH4+ assimilation, where the enzyme GS facilitates this conversion. Following GS action, one of the amino groups from glutamine is transferred to 2-oxoglutarate (2-OG) to synthesize glutamate, and this reaction is catalyzed by GOGAT (Figure 3). The coordinated action of GS and GOGAT is responsible for these interconnected processes. Experimental studies involving reverse genetic analysis have demonstrated that the absence of genes encoding NH4+-responsive GS/GOGAT isoenzymes reduces NH4+ assimilation, particularly in the roots, resulting in impaired growth. These findings suggest that the NH4+-responsive forms of GS/GOGAT have a central role in the primary assimilation of NH4+ within the roots. As the expression of these isoenzyme-encoding genes increases with NH4+ availability, it is plausible to infer that plants possess a regulatory transcriptional network that modulates their gene expression in response to NH4+ availability.

Plant GS (EC6.3.1.2), a highly conserved enzyme, catalyzes glutamine synthesis from glutamate and ammonium ions, conserving the N atom in an organic molecule. This conversion is fueled by the energy released through the hydrolysis of ATP to ADP, making GS a crucial component of N metabolism across different life forms. In addition, the GOGAT in synthesizing glutamate involves transferring the amide amino group of glutamine to 2-OG through a reductant-driven process, yielding two glutamate molecules. Within the realm of plants, this enzyme exists in two distinct variants: one produces the ferredoxin (Fd) as the electron donor (EC 1.4.7.1), while the other employs NADH as the electron donor (EC 1.4.1.14). The Fd-dependent form of the enzyme is notably abundant within the chloroplasts of photosynthetic tissues. In these regions, it utilizes light energy directly to serve as a reductant source. Figure 3 provides an overview of the N cycle, transport, and utilization in plants.

Conversely, the NADH-dependent variant, also present within plastids, is primarily concentrated in non-photosynthesizing cells. The reductant is sourced from the pentose phosphate pathway. Notably, the expression of the Fd- and NADH-dependent forms of glutamate synthase seems to exhibit dissimilar patterns across distinct plant tissues. It has been suggested that in most plants, two genes encode each form of GOGAT.

4.2. Isoforms of GS and GOGAT

According to subcellular distribution patterns, investigations into plant GS enzymes have identified two main categories. The initial group, designated as GS1, is primarily situated within the confines of the cytoplasm. Conversely, the second group, GS2, is predominantly found within plastid structures. Various multigenic families are responsible for encoding multiple isoforms of GS1, whereas the plastidial GS2 arises from a limited number of nuclear genes. Typically, GS2 is linked to the process of (re)assimilation of NH4+ in leaves, whereas GS1 is associated with recycling N within the plant. However, the relative activity levels exhibited by GS1 and GS2 are subject to variation based on species, specific plant organs, N sources, developmental stages, and prevailing environmental conditions, including abiotic stress factors. This variability underscores the intricate involvement of these isozymes, reflecting a multifaceted role. In wheat research, a comprehensive exploration has unveiled four distinct variations of GS genes: GS1, GS2, GSr, and GSe. Employing advanced high-performance liquid chromatography (HPLC) methods, ref. successfully separated GS and discovered two subtypes in leaves: cytoplasmic subtype GS1 and chloroplast subtype GS2. Both subtypes’ content and activity fluctuate in tandem with developmental processes. The subcellular localizations of GSr and GSe align with the cytoplasmic compartment. In addition, the advancement of genome sequencing has unveiled various isoenzymes of GS/GOGAT present in plants. Among these, specific isoenzymes such as GS1,2 and NADH-GOGAT1 from rice as well as GLN1,2 and NADH-GOGAT (GLT1) from Arabidopsis are expressed in plant roots in response to NH4+ supply. These isoenzymes’ expression at transcript and protein levels highlights their specificity in NH4+ assimilation.

4.3. Nitrogen Metabolism in Microalgae

Microalgae are unicellular aquatic microorganisms with sizes between 1 µm and 2 mm. They can generate biomass using light and carbon dioxide through the photosynthetic process. Due to their remarkable tolerance for environmental stress, they can be grown in various environmental conditions, including freshwater, saltwater, or wastewater. After C, N is quantitatively the most significant element contributing to the dry matter of microalgal cells, accounting for between 1 and 10% of their dry weight. C and N metabolism are related in microalgae since they share the following: (a) C provided directly by respiration of CO2 or assimilated organic C and (b) the energy obtained by the TCA cycle and mitochondrial electron transport chain. To create the amino acids glutamate, glutamine, and aspartate, the primary absorption of inorganic N (NH4+) needs carbon skeletons in the form of keto acids (2-OG and oxaloacetate) as well as energy in the form of ATP and NADPH. There are four fundamental phases in the NO3− assimilation mechanism in eukaryotic microalgae and plant leaves: NAD (P) H-nitrate reductase catalyzes NO3− reduction into NO2− in the cytosol of cells, which is required for (a) NO3− transport into the green cell, and (b) NO3− reduction into NO2− in the cytosol of cells. In addition, in Cyanobacteria, ferredoxin-nitrate reductase catalyzes the reduction of NO3− to NO2−, (c) NO2− transport into the chloroplast of cells via NAR1 family and its subsequent reduction into NH4+ in a six-electron reaction catalyzed by ferredoxin-nitrite reductase, and (d) the process mediated by the ATP-dependent GS- GOGAT cycle, which results in the formation of L-glutamate by the incorporation of NH4+ into the carbon skeleton of 2-OG. Microalgae mostly use the GS-GOGAT pathway to assimilate NH4+ as well as via the amination of 2-OG catalyzed by NAD(P)H glutamate dehydrogenase [NAD(P)H-GDH.

5. The Role of Nitrogen in Plant Response to Abiotic Stress

Plants must endure abiotic stressors, including drought, salt, and severe temperatures, because they are immobile. The spread of plants is significantly constrained by these stresses, which also change plant growth and development and lower agricultural output. Since N is considered the most crucial nutrient for plant growth quantitatively, plants have developed effective methods to control N levels in response to complicated stresses.

5.1. Nitrogen and Drought Stress

Drought stress significantly damages crops worldwide, particularly in arid and semi-arid regions. Hence, plants have evolved to adapt their resilience against drought via avoidance (stomata adjustment to reduce transpiration rate) and tolerance (antioxidants and osmolytes production) of drought (Figure 4). Remarkably, the concentrations of NO3− and NH4+ have impacts on plants’ underwater limitations. The application of NH4+ alleviates the effect of drought on plant development, while NO3− has the opposite impact. In that sense, N fertilizers might enhance the physiological responses of plants under drought stress by elevating the concentrations of N and chlorophyll and promoting PSII photochemical activity. It has been shown that adding N (NH4NO3) improved water-use efficiency (WUE) by increasing plant dry mass and decreasing water loss.

Additionally, N may lessen drought’s inhibitory effects on photosynthesis and prevent C starvation. On the other hand, low N availability can make plants more sensitive to dehydration and cause protein degradation, which lowers the amount of N-containing osmolytes like proline. N supply enhances osmotic adjustment and the activity of antioxidant enzymes via proline accumulation induction to mitigate the physiological harm from drought stress. However, the N effects depend on the ionic form of N supplied. NH4+ is essential in promoting plant drought tolerance by increasing root numbers (surface area) and subsequently improving and facilitating water uptake. Also, decreased aerenchyma development may contribute to NH4+-promoted drought tolerance. The influence of NO3− on drought responses is linked to NO3− transport/assimilation in plants. Numerous NO3− transport/assimilation-related genes, such as NRT2.5, GS, and GOGAT, are repressed in response to drought stress. Interestingly, a high N concentration enhances root growth and photosynthetic traits, alleviates drought stress via stomatal conductance, and increases the photosynthetic rate. Plant nutrient management is crucial to promote abiotic stress tolerance in cotton plants. The use of nitrogen fertilizer either in the soil or foliar spray can decrease the toxicity of reactive oxygen species (ROS) by increasing the peroxidase (POD), catalase (CAT), and superoxide dismutase (SOD) levels in plant cells. These enzymes improve the photosynthetic rate, promoting stress tolerance by scavenging ROS and reducing photooxidation to preserve chloroplast membrane integrity. Applying nitrogen fertilizers (N60 kg.ha−1) can increase the tolerance of drought stress in wheat plants combined with the highest yield.

Under drought stress, GS1 and GS2 exhibited differential expression among different plant organs and species, with a prominent increase of GS1 in roots and leaves, while the total GS activity showed a decrease in potato root. A comparative proteomic analysis under drought stress conditions in wheat was studied. The researchers found that drought-tolerant wheat varieties exhibited a significant upregulation of GS2 expression, indicating its potential role in enhancing drought tolerance. Similar results were obtained by Nagy et al., who explored GS activity in wheat leaves and observed that older leaves had lower GS activity than flag leaves in drought-tolerant and sensitive varieties under normal conditions. During drought stress, GS activity in sensitive varieties’ flag leaves decreased compared to tolerant varieties. This suggested that GS activity could indicate drought stress sensitivity and tolerance, with elevated cytoplasmic-to-plastid GS ratio and premature senescence as markers. Exposure to heavy metals such as cadmium, molybdenum, copper, and aluminum led to the down-regulation of GS and decreased its total activity in different plant species, including rice, soybean, tomato, and tea.

Like all organisms, plants must respond to various extreme environmental cues by responding to various internal signals. NO is a small lipophilic molecule that diffuses through plant cell membranes as active signals to thrive and survive under those conditions. In addition, NO is a relevant signal molecule in many plant processes. Several pathways for NO production, either oxidative in the presence of O2 or reduction from NO2−, have been described in plants. NO regulates gene expression, modulates enzyme activities, and acts as a metabolic intermediate in energy regeneration. However, the mechanisms and proteins responsible for its synthesis remain complex, with many unresolved questions.

Moreover, several studies showed the involvement of NO in reproductive processes, control of development, and the regulation of physiological responses such as stomatal closure. NO also regulates the expression of several genes involved in the synthesis of and response to pathogen attacks as well as reproductive mechanisms that operate during pollen recognition by the stigma. NO is a high redox to reactive N species (RNS) formed in living cells under normal and stressful biotic and abiotic conditions, and NO also functions as a detoxifier and lessens adverse effects when a plant’s ROS content reaches dangerous levels. It was reported that the synthesis of NO increased in plant cells under drought stress conditions, and it has been suggested that this may help enhance plants’ acclimation responses to cope with water limitation. Drought stress enhanced NO synthesis in beat, pea, tobacco, cucumber, grapevine, and rice plants; likewise, the activity of NO synthase (NOS)-like and NO release rate increased under dehydration conditions. NO has a vital role in respiratory function, called pathways of electron transport in mitochondria; it adjusts ROS as well as activates defense strategies via promoted antioxidant formation in plants under various abiotic stresses. NO plays a crucial role in mediating hormonal activities and influencing gene expression as well as protein activity during sensing and signaling processes that enable plants to survive under environmental stressors. The exogenous supply of NO increased drought tolerance in various plants such as maize, rice, and cucumber due to the increase of antioxidant defense, which acts to scavenge ROS, improve cellular membrane stability, and preserve photosynthesis as well as water status.

Drought induces the synthesis of abscisic acid (ABA), which regulates several critical plant processes like seed germination and stomatal conductance, with the help of signaling molecules like NO and ROS. Interestingly, NO acts as a critical signaling molecule promoting ABA-induced stomatal closure in case of hyperosmotic stress. The plants’ main enzyme responsible for NO production is the molybdoenzyme NR, which contains two subunits, each bearing three prosthetic groups: FAD, heme b557, and molybdenum. In an NR subunit, molybdenum is bound to a tricyclic pyranopterin and chelated by a dithiolene named the molybdenum cofactor (Moco). Amongst the reducing sources of NO in plants, NO2− can be reduced to NO by a family of five molybdenum (Mo)-containing enzymes, which include NR, xanthine oxidase reductase (XOR), aldehyde oxidase (AO), sulfite oxidase (SO), and mARC.

In a study by Berger et al. (2020) three NR genes (MtNR1, MtNR2, and MtNR3) were discovered in the genome of Medicago truncatula, and they study addressed their expression, activity, and potential involvement in NO production during the symbiosis between M. truncatula and Sinorhizobium meliloti. These results revealed that MtNR1 and MtNR2 gene expression and activity are correlated with NO production throughout the symbiotic process and that MtNR1 is mainly involved in NO production in mature nodules. However, when plants were grown in soil containing NH4+, the functional role of NR was eliminated. Moreover, NO can alleviate the negative impact of ROS through lipid peroxidation, enhancing photosynthesis as well as the expression of antioxidant enzymes via mitogen-activated protein kinase (MAPK) and other signaling pathways. It was found that exogenous application of NO caused a significant increase in seedling growth of cucumber by decreasing seedlings injury and enhancing the antioxidant activities and chlorophyll and free proline contents under stress conditions. Moreover, an exogenous supply of NO in wheat markedly preserved high relative water content via decreasing transpiration and increasing K content and mitigated cell membrane damage. Figure 4 explains how NO can work as a signaling molecule that can protect plants from drought stress by modulating various physiological and biochemical processes.

5.2. Nitrogen and Salinity Stress

Salinity stress has a destructive impact on the metabolism and productivity of crops worldwide, resulting in the remarkable conversion of agricultural arable soil into unproductive land. The high level of salt ions in agricultural soil may be harmful to plants, causing salt stress, where sodium (Na+) and chlorine (Cl−) concentrations increase in saline soil and affect the natural physiological process in plants. N application is an efficient way to improve plants under saline conditions. N is a necessary nutrient that mediates various molecular, physiological, and cellular responses essential to plant survival as well as different signal transduction pathways linked to plant defense systems and salt-stress tolerance. Moreover, salt stress affects ammonification and nitrification in the soil because Cl− competes with NO3−, and NH4+ competes with Na+ and creates ion toxicities and inequalities, which can limit the processes of N uptake, transport, and assimilation. Salt stress has inhibited various enzyme activities that contribute to N assimilation in plants such as maize, tomato, rice, cowpea, and mung bean, reducing N uptake and utilization.

Interestingly, it was reported that the interaction between salt stress and N had an impact on plant growth, and the effect of N on salt-exposed plants was dependent on the N rate. In the case of maize, excessive N treatment can successfully mitigate the deleterious effects of salt stress. Similar findings in tomatoes showed that supra-optimum N was better than optimum N in reducing salt stress. Under salinity stress, the application of N significantly increased both N and K uptake and decreased Na accumulation in wheat seedlings. Furthermore, N application can protect wheat seedlings in saline conditions via the upregulation of osmolytes, antioxidant system, and secondary metabolite accumulation. Even though excessive N application exacerbated the negative effect of salt stress on wheat and rice, moderate N supply could ameliorate the deleterious effect of salt stress. Therefore, applying an appropriate N rate is the key to mitigating the adverse effects caused by salt stress.

On the contrary, it was reported that the application of NO3− resulted in an increase of Na+ uptake and loading of the sodium into the xylem, increasing root inhibition caused by the salinity. Na+ and NO3− co-transportation was suggested based on NO3−-dependent Na+ uptake data at various Na+ concentrations. Similarly, NO3−-dependent transport mechanisms in salt-stress environments encourage Na+ ion uptake and loading into the xylem, which may represent a main pathway for Na+ accumulation in Arabidopsis shoots.

It is worth noting here that GS isoforms imply a multifaceted role for GS in navigating through various dimensions of abiotic stress tolerance. For instance, in rice plants exposed to salinity stress, GS1 increased in the root at the seedling stage and old leaves with no change in young leaves, while GS2 was down-regulated in old and young leaves.

It has been demonstrated that NO can act as a signal to induce the resistance of salt stress by raising the K:Na ratio in plant cells, which is based on an increase in the H1-ATPase activity of plasma membrane in calluses from maize seedlings, reed (Phragmites communis) plants, and poplar trees. Exogenous NO can improve salt tolerance under salinity conditions by stimulating the K+/Na+ ratio, the Na+/H+ antiport in the tonoplast, and proton pump activity. Under salt stress, NO improves the absorption and translocation of several macro- and micronutrients, including K, Fe, Mg, and Zn, which improve respiration and chlorophyll biosynthesis. The uptake of mineral nutritional elements was improved due to NO application, thereby diminishing the harmful effect of high salinity on strawberry plants. The exogenous NO supply can protect chickpea plants from salt stress-induced oxidative harm by promoting the biosynthesis of photosynthetic pigment and antioxidant enzymes osmolyte accumulation, thereby improving plant development under saline stress. Furthermore, under salt stress conditions, exogenous NO treatment enhanced wheat seed germination by converting starch to sugars, where the application of NO led to an elevation in the effectiveness of α- and β-amylase enzymes. Under salt stress, NO treatment boosted plant development and demonstrated a protective ability against salinity-triggered oxidative stress by increasing the effectiveness of POD, SOD, and CAT enzymes in Triticum aestivum.

6. Hazardous Effects of N Fertilizers

Nutrient deficiency and/or availability are crucial to plant production. One of the severe issues the world faces is the adverse effects of fertilizers on the environment. Farmers have been using fertilizers since ancient times, but long-term use has affected soil fertility adversely. Due to over-fertilization in many agricultural areas, environmental pollution was established. N pollution in groundwater is a severe problem worldwide, affecting animal and human health; in addition, NO3− leaching decreases N availability for crops and can cause water pollution. The improper application of fertilizers damages the farmlands, leading to heavy metal contamination and soil erosion. While ammonium sulfate causes soil pollution, nitrogen oxides and NH4+ cause air pollution, and NO3− is the final breakdown product accumulating in groundwater. Since NO3− levels above 10 mg L−1 N cause oxygen debt in the blood, the U.S. Environmental Protection Agency set human drinking water to maintain nitrate levels below a maximum contaminant level (MCL) at a critical value of 10 mg L−1 N. In Argentina, the national food law set the limit for human consumption at 11.3 mg L−1 N. Consumption of NO3−-rich water has been associated with methemoglobinemia (blue child syndrome) and different forms of cancer. Furthermore, diabetes, adverse reproductive outcomes (especially neural tube defects), and thyroid conditions are related to NO3−-contaminated drinking water.

A notable study by Maghanga et al. (2013) was conducted to monitor changes in surface water NO3− levels in ten rivers within a Kenyan tea plantation for three years. Water samples were obtained before and after fertilizer application in 2004, 2005, and 2006. For three years, there was no established trend between surface water nitrate levels and the time of fertilizer applications. However, the highest nitrate–nitrogen levels were in thr river Temochewa during the first fertilizer applications (4.9 mg L−1 to 8.2 mg L−1). Furthermore, fertilizer application increased NO3− levels, and the study indicated that initial nitrate–nitrogen levels in most rivers were high, causing surface water contamination.

7. Promoting N-Fixation through Diazotrophic Microbiota

Rapid human population growth and declining agricultural soils intensify resource competition, affecting human food hunger and sustainable agriculture systems worldwide. Hence, developing alternative strategies to increase the current food demand is an inevitable need. Accordingly, crop productivity is the pillar of nutritional food security and heavily depends on applying N fertilizers. Applying N fertilizers consumes vast amounts of energy and causes excessive harm to the environment and health, expensive costs, soil fertility reduction, and other negative consequences. Therefore, increasing plant NUE is essential for developing sustainable agriculture. Ultimately, N fixation has multiple benefits through improving the yields of landraces that often receive little attention from breeders and empowering farmers using a sustainable and less expensive strategy. Additionally, it reduces the need for fossil fuels requiring synthetic N fertilizers. Overall, the symbiotic relationship between leguminous plants and Rhizobia is a powerful adaptation strategy that enables these plants to reduce their dependence on synthetic fertilizers and thrive in N-deficient soils. Improving the microbial activities that boost plant growth is necessary to increase food production. In that sense, several beneficial mutualistic microbes have been discovered; however, their dependable utilization as biofertilizers in soil conditions is still challenged. Thus, improving the microbial inoculants and exploring their diversity performance in the external habitat could lead to understanding the knowledge gap between the microbial performance in vitro and the external habitat.

Bacterial endophytes associated with crops, i.e., corn and wheat, can produce various sugar compositions, such as arabinose, to stimulate the production of a thick gelatinous layer (mucilage) around the root systems of these crops. This layer allows crops to better uptake fertilizer by forming a continuous boundary layer between roots and soil particles, protecting the root system from drought and supporting other microbes (Figure 5). Most interestingly, mucilage structures associated with complex microbiota contribute to nutrient acquisition and play a vital role in plant growth and defense.

Improving the sustainability of biological nitrogen fixation (BNF) to cereal crops is a future reason to enhance crop productivity, representing a significant breakthrough in N-fixation research. To achieve this goal, engineering key regulators of BNF by activating the symbiotic signaling pathway between diazotrophic bacteria and cereal crops is a promising strategy for sustainable development.

Fortunately, establishing new omics techniques such as metabolomics, metagenomics, metatranscriptomics, and metaproteomics will help in-depth research and identify the rhizospheric microbes using next-generation sequencing approaches. Accordingly, manipulating the promising microbes that replace synthetic fertilizers is a solution. Rhizobium–plant symbiosis is now becoming challenging with diverse genetic modifiers of the symbiosis relationship not only in the genomic variability of partners but also in the soil microbiota. In that sense, there is a future direction to improve the specificity of the partnership during symbiotic colonization using different methods (e.g., axenic cultures) by eliminating other microbes. Therefore, identifying and developing bioinoculant strains and genotypes by adapting and evolving elite strains in both lab-scale analyses and trials to open-field applications to increase crop yield becomes crucial. Figure 5 shows how microbes can help plants increase their N availability by colonizing their roots, releasing exudates, and enhancing nutrient uptake.

8. Microbial Alternatives to Synthetic Fertilizers

Fertile soil is a primary component in the backbone of sustainable agriculture; however, land degradation and rapid desertification cause an estimated global loss of 24 billion tons of fertile arable land. Noticeably, eutrophication and decline of soil fertility result from massive application of chemical fertilizers, causing significant environmental concern. Moreover, with the limited N resources and increasing agricultural demands for N supplies, there is a need to discover diverse mutualistic interactions between plant roots and rhizosphere microbiome (Figure 5).

Soils are habitats for diverse microorganisms that are either harmful (pathogenic) or beneficial (symbiotic) to plants, comprising organic substances, water, and nutrients. Food production requires essential nutrients, metabolites, and water that the soil provides to plants. Besides the nutrient resources in the soil, plants attract microbes via root exudates, root border cells, and mucilage formation that serve as food for the rhizosphere microbiome and their assembly. Interestingly, soil contains an extensive reservoir of microorganisms (1 × 109 microbial cells g−1 dry soil) and microbial diversity (1 × 105 microbial species g−1 dry soil). Rhizospheric microbes are crucial in diverse biological processes, including soil structure, climate regulation, disease control, and organic matter decomposition. They also induce plant growth via nutrient uptake, cycling water availability, and the formation of stable aggregates to reduce the risks of soil erosion. In addition, these microbes can improve plant health and serve as biocontrol agents against plant pathogens as well as biofertilizers (i.e., Rhizobium, Azotobacter, and Bacillus subtilis). Accordingly, elucidating the functional role of these microbial communities has garnered enormous efforts from botanical scientists over two decades to eliminate hunger and improve soil fertility by reducing synthetic fertilizers. Fortunately, legume plants represent a solution, including fava beans, soybeans, lentils, and cowpea. They recruit symbiotic Rhizobia from the soil into nodules BNF. BNF is an eco-friendly and manageable strategy by harnessing nitrogen-fixing endophytic and free-living rhizobacteria to increase N levels in agricultural land by converting N2 to the fixed form, NH3. The biologically fixed amounts of atmospheric N into NH3 inside root nodules by legumes account for about 65% of N utilization in global agriculture via N-fixing Rhizobia. Accordingly, this process allows legume plants to grow well in N-deficient soils, eliminating N fertilizer application. The inoculation of Rhizobia strains (native or commercial) was studied on the growth and nodulation of three cowpea (Vigna unguiculata) genotypes in semiarid regions of Kenya. Field trials were performed in a randomized complete block design with three replications, and the symbiotic efficiency (SE) of Rhizobium isolates was evaluated. In the field, Rhizobia inoculation significantly increased nodulation and shoot DW compared to the uninoculated controls. Interestingly, Rhizobia inoculation significantly increased yields, whereas inoculation with native isolates recorded a 22.7% increase compared to uninoculated control in the first season and a 28.6% increase in yield in the second season. This study concluded that the efficient native Rhizobia in smallholder farms has promising potential to improve cowpea yield under a changing climate. In particular, R. tropici clone H53, Mesorhizobium sp. WSM3874, and R. pusense strain Nak353 showed more superiority in all the tested parameters. A recent study revealed that plant exudates play a role in the development of bacterial biofilms (multicellular communities of microorganisms) in the soil, and creating microaerophilic conditions improved the nitrogen-fixing efficacy of diazotrophic bacteria in the soil and exhibited enhanced flavonoid compounds. This study also hypothesized that manipulating flavonoid synthetic pathways leads to the induction of BNF in cereals through biofilm formation in soil diazotrophs. N-limitation induces the production of exopolysaccharides (EPS), which may function as a barrier to block excess oxygen in the air and provide a suitable microaerobic condition for bacteria inside the EPS to fix N. A large amount of EPS was reported in response to N limitation by cellulolytic bacteria (e.g., Bacillus, Pedobacter, Chryseobacterium, and Flavobacterium.

Applying the symbiotic association between Rhizobia and leguminous plants is a significant research area to reduce the dependency on chemical fertilizers. Bradyrhizobium, Azospirillum, and Rhizobium had high plant-growth-promoting capacity and N-fixing efficiency, indicating abundant functional microbial resources in extreme soil environments. On the other hand, inoculation of Azospirillum, Burkholderia, and other N-fixing bacteria had little success in the field. Thus, developing a compatible host for the indigenous N-fixing bacteria is critical for colonizing the host and improving nitrogenase activity.

9. Genomic Approaches for Improving N-Fixation

The identification of new genes involved in N fixation, investigation of the genome organization of N-fixing species, and the discovery of novel diazotrophic species have all been made possible by plant and microbial genomics. The primary genes responsible for N fixation encode nitrogenase enzymes, which play a crucial role in converting N2 into NH4+. These enzymes are classified into several types based on their structural, evolutionary, and functional characteristics. The first three forms of nitrogenase share similarities in their architecture, evolutionary origins, and mechanisms. However, they differ in the type of metal they use as a cofactor, either vanadium or molybdenum, or they may rely solely on iron as their cofactor. In contrast, the fourth type of nitrogenase operates differently; it utilizes a molybdopterin-containing system and depends on the presence of superoxide. This distinct form of nitrogenase was initially identified in a specific species called Streptomyces thermoautotrophicus. The complete genomic sequence of S. thermoautotrophicus and the precise amino acid sequence of this fourth type of nitrogenase have not been determined. The fact that the inventory and distribution of species encoding for the fourth kind of nitrogenase are unknown highlights how few diazotrophs’ genomes have been sequenced. A. vinelandii is one of the extensively researched diazotroph with sequenced genomes that can fix N in an aerobic culture. It has served as a model for N-fixation for decades due to several factors, including (i) its genetic adaptability; (ii) its capacity to fix N in aerobic growth conditions; (iii) its nutritional flexibility, which is demonstrated by its capacity to fix N via three different pathways; and (iv) its capacity to adapt its metabolism to a variety of nutrients and media additives; and (v) the genome’s comprehensive sequencing and manual editing. A. vinelandii has a single circular chromosome with 5,365,318 base pairs in its genome, which is expected to code for 5051 proteins. The diazotroph Pseudomonas stutzeri is the closest companion of A. vinelandii with a sequenced genome. Moreover, P. stutzeri grows aerobically and can only fix N in microaerobic conditions. Alternatively, in ambient oxygen concentrations (20% O2), A. vinelandii can catalyze N fixation in an oxygen-sensitive process. Its respiratory protection system makes it feasible to undertake two incompatible biological processes, oxidative phosphorylation and N fixation, concurrently. A. vinelandii may modify its respiration rate to maintain a low amount of cytoplasmic oxygen while N-fixing is taking place. Five terminal oxidases were found in the genome, along with additional NADH oxidoreductases and other respiratory complexes that provide electrons to terminal oxidases, raising oxygen consumption. Some of these genes, such as cydAB I, have been linked to respiratory protection and are crucial for aerobic N-fixation. The discovery of novel N-fixing organisms and the quantity and kind of nitrogenases coding in these species may also be targeted by genome sequencing utilizing NifD sequences as queries. The genomes of diazotrophs also encode for other proteins comparable to NifD in addition to the Nif/Vnf/AnfD sequences.

10. Conclusions and Future Perspectives

N is an essential macronutrient required for the growth and development of all living organisms. N uptake, transport, assimilation, and metabolism are finely tuned in plants. Additionally, N serves as a key mediator in plant–microbe interactions, fostering beneficial relationships through symbiotic partnerships that enhance nutrient availability. Moreover, N plays a pivotal role in a plant’s response to various environmental stresses, such as drought, salinity, and disease. Understanding N dynamics is critical for optimizing crop yields and sustainability, as it impacts the health and productivity of individual plants and the entire ecosystem and food security.

Consequently, harnessing N in agriculture addresses the challenges of feeding a growing global population while minimizing the environmental footprint of farming practices. In this sense, improving NUE by plants is a promising strategy to reduce the negative impact of using synthetic fertilizers. BNF is an eco-friendly and manageable strategy to increase N levels in agricultural land. Thus, improving BNF by genome editing could replace synthetic fertilizers to reduce dependency on chemical fertilizers. In addition, discovering new plant microbiomes and understanding their structure, abundance, and diversity will open a new avenue to better understanding the agricultural scenario to sustain global food security under limited N conditions. Despite understanding the role of N in plant biology, the precise molecular mechanisms governing N sensing in plants are still unclear, which could lead to the development of more efficient N-use strategies.

Furthermore, investigating the effects of climate change on N availability and its environmental impact becomes increasingly crucial. For example, arctic land areas are expected to warm, resulting in significant permafrost thawing and significant alterations in ecological functioning. With thawing, a massive pool of immobile C trapped in permafrost becomes available for breakdown and remobilization, resulting in CO2 and methane (CH4) emissions. The gaseous C leak from thawing permafrost is being extensively researched in order to determine the extent of the permafrost–carbon feedback to the climate. Permafrost soils, on the other hand, are huge N reservoirs, with an approximate amount of 67 billion tons of total N in the top 3 m. Organically bound N is mineralized upon thawing, resulting in the release of NH4+ and NO3− and driving nitrification and denitrification, the two primary processes producing greenhouse gas in soils, and these microorganisms can convert to harmful NO or N2 gas. In addition to methane and carbon dioxide fluxes, warming arctic soils release N, which can increase N2O emissions, another potent greenhouse gas. The significance and amount of N2O released from arctic soils are unknown. There is a crucial need for a combination of in situ field investigations, laboratory experiments, and cutting-edge metagenomic techniques to better understand the role of microorganisms in climate change and seasonal changes in bacteria.

References 327

  1. 1.

    Lal R. Restoring Soil Quality to Mitigate Soil Degradation. Sustainability. 2015;7:5875-5895.

    Not in our library10.3390/su7055875

  2. 2.
    Soil and the intensification of agriculture for global food security · 2019
    Not checked
  3. 3.

    Pereira P., Bogunovic I., Muñoz-Rojas M., Brevik E.C. Soil ecosystem services, sustainability, valuation and management. Curr. Opin. Environ. Sci. Health. 2018;5:7-13.

    Not in our library10.1016/j.coesh.2017.12.003

  4. 4.

    Pimentel D., Burgess M. Soil Erosion Threatens Food Production. Agriculture. 2013;3:443-463.

    Not in our library10.3390/agriculture3030443

  5. 5.

    Díaz-Rodríguez A.M., Salcedo Gastelum L.A., Félix Pablos C.M., Parra-Cota F.I., Santoyo G., Puente M.L., Bhattacharya D., Mukherjee J., de los Santos-Villalobos S. The Current and Future Role of Microbial Culture Collections in Food Security Worldwide. Front. Sustain. Food Syst. 2021;4:614739.

    Not in our library10.3389/fsufs.2020.614739

  6. 6.

    Arora N.K., Khare E., Maheshwari D.K., Maheshwari D.K. Plant Growth Promoting Rhizobacteria: Constraints in Bioformulation, Commercialization, and Future Strategies. Plant Growth and Health Promoting Bacteria. 2011.

    Not in our library

  7. 7.

    Hirel B., Tétu T., Lea P.J., Dubois F. Improving Nitrogen Use Efficiency in Crops for Sustainable Agriculture. Sustainability. 2011;3:1452-1485.

    Not in our library10.3390/su3091452

  8. 8.

    Akinnifesi F.K., Ajayi O.C., Sileshi G., Chirwa P.W., Chianu J. Fertiliser trees for sustainable food security in the maize-based production systems of East and Southern Africa. A review. Agron. Sustain. Dev. 2010;30:615-629.

    Not in our library10.1051/agro/2009058

  9. 9.

    Spiertz H. Avenues to meet food security. The role of agronomy on solving complexity in food production and resource use. Eur. J. Agron. 2012;43:1-8.

    Not in our library10.1016/j.eja.2012.04.004

  10. 10.

    Azadi H., Ho P. Genetically modified and organic crops in developing countries: A review of options for food security. Biotechnol. Adv. 2010;28:160-168.

    Not in our library10.1016/j.biotechadv.2009.11.003

  11. 11.

    Singh J.S., Gupta V.K. Soil microbial biomass: A key soil driver in management of ecosystem functioning. Sci. Total Environ. 2018;634:497-500.

    Not in our library10.1016/j.scitotenv.2018.03.373

  12. 12.

    Shrivastav P., Prasad M., Singh T.B., Yadav A., Goyal D., Ali A., Dantu P.K. Role of nutrients in plant growth and development. Contaminants in Agriculture: Sources, Impacts and Management. 2020;43-59.

    Not in our library

  13. 13.

    Zhang S., Liu Y., Du M., Shou G., Wang Z., Xu G. Nitrogen as a regulator for flowering time in plant. Plant Soil. 2022;480:1-29.

    Not in our library10.1007/s11104-022-05608-w

  14. 14.

    Ye J.Y., Tian W.H., Jin C.W. Nitrogen in plants: From nutrition to the modulation of abiotic stress adaptation. Stress Biol. 2022;2:4.

    Not in our library10.1007/s44154-021-00030-1

  15. 15.

    Hu B., Chu C. Nitrogen–phosphorus interplay: Old story with molecular tale. New Phytol. 2020;225:1455-1460.

    Not in our library10.1111/nph.16102

  16. 16.
    Effects of Potassium Levels on Plant Growth, Accumulation and Distribution of Carbon, and Nitrate Metabolism in Apple Dwarf Rootstock Seedlings · 2020
    No notice
  17. 17.

    Li H., Yu M., Du X.-Q., Wang Z.-F., Wu W.-H., Quintero F.J., Jin X.-H., Li H.-D., Wang Y. NRT1.5/NPF7.3 functions as a proton-coupled H+/K+ antiporter for K+ loading into the xylem in Arabidopsis. Plant Cell. 2017;29:2016-2026.

    Not in our library10.1105/tpc.16.00972

  18. 18.

    Kosegarten H.U., Hoffmann B., Mengel K. Apoplastic pH and Fe3+ reduction in intact sunflower leaves. Plant Physiol. 1999;121:1069-1079.

    Not in our library10.1104/pp.121.4.1069

  19. 19.

    Zhao T., LING H.Q. Effects of pH and nitrogen forms on expression profiles of genes involved in iron homeostasis in tomato. Plant Cell Environ. 2007;30:518-527.

    Not in our library10.1111/j.1365-3040.2007.01638.x

  20. 20.

    Scherer H.W. Impact of sulfur on N2 fixation of legumes. Sulfur Assimilation and Abiotic Stress in Plants. 2008;43-54.

    Not in our library

  21. 21.

    Clarkson D.T., Saker L.R., Purves J.V. Depression of nitrate and ammonium transport in barley plants with diminished sulphate status. Evidence of co-regulation of nitrogen and sulphate intake. J. Exp. Bot. 1989;40:953-963.

    Not in our library10.1093/jxb/40.9.953

  22. 22.

    Rietra R.P., Heinen M., Dimkpa C.O., Bindraban P.S. Effects of nutrient antagonism and synergism on yield and fertilizer use efficiency. Commun. Soil Sci. Plant Anal. 2017;48:1895-1920.

    Not in our library10.1080/00103624.2017.1407429

  23. 23.

    Saud S., Fahad S., Yajun C., Ihsan M.Z., Hammad H.M., Nasim W., Jr A., Arif M., Alharby H. Effects of nitrogen supply on water stress and recovery mechanisms in Kentucky bluegrass plants. Front. Plant Sci. 2017;8:983.

    Not in our library10.3389/fpls.2017.00983

  24. 24.

    Masclaux-Daubresse C., Daniel-Vedele F., Dechorgnat J., Chardon F., Gaufichon L., Suzuki A. Nitrogen uptake, assimilation and remobilization in plants: Challenges for sustainable and productive agriculture. Ann. Bot. 2010;105:1141-1157.

    Not in our library10.1093/aob/mcq028

  25. 25.

    Sandhu N., Sethi M., Kumar A., Dang D., Singh J., Chhuneja P. Biochemical and genetic approaches improving nitrogen use efficiency in cereal crops: A review. Front. Plant Sci. 2021;12:657629.

    Not in our library10.3389/fpls.2021.657629

  26. 26.

    Delin S., Stenberg B., Nyberg A., Brohede L. Potential methods for estimating nitrogen fertilizer value of organic residues. Soil Use Manag. 2012;28:283-291.

    Not in our library10.1111/j.1475-2743.2012.00417.x

  27. 27.

    Yao X., Nie J., Bai R., Sui X. Amino acid transporters in plants: Identification and function. Plants. 2020;9:972.

    Not in our library10.3390/plants9080972

  28. 28.

    Li F., Dong C., Yang T., Bao S., Fang W., Lucas W.J., Zhang Z. The tea plant CsLHT1 and CsLHT6 transporters take up amino acids, as a nitrogen source, from the soil of organic tea plantations. Hortic. Res. 2021;8:178.

    Not in our library10.1038/s41438-021-00615-x

  29. 29.

    Feng H., Fan X., Miller A.J., Xu G. Plant nitrogen uptake and assimilation: Regulation of cellular pH homeostasis. J. Exp. Bot. 2020;71:4380-4392.

    Not in our library10.1093/jxb/eraa150

  30. 30.

    Forde B.G. Nitrate transporters in plants: Structure, function and regulation. Biochim. Et Biophys. Acta (BBA)-Biomembr. 2000;1465:219-235.

    Not in our library10.1016/s0005-2736(00)00140-1

  31. 31.

    Iqbal A., Qiang D., Zhun W., Xiangru W., Huiping G., Hengheng Z., Nianchang P., Xiling Z., Meizhen S. Growth and nitrogen metabolism are associated with nitrogen-use efficiency in cotton genotypes. Plant Physiol. Biochem. 2020;149:61-74.

    Not in our library10.1016/j.plaphy.2020.02.002

  32. 32.

    Léran S., Varala K., Boyer J.-C., Chiurazzi M., Crawford N., Daniel-Vedele F., David L., Dickstein R., Fernandez E., Forde B. A unified nomenclature of nitrate transporter 1/peptide transporter family members in plants. Trends Plant Sci. 2014;19:5-9.

    Not in our library10.1016/j.tplants.2013.08.008

  33. 33.

    Wang Y.-Y., Hsu P.-K., Tsay Y.-F. Uptake, allocation and signaling of nitrate. Trends Plant Sci. 2012;17:458-467.

    Not in our library10.1016/j.tplants.2012.04.006

  34. 34.

    Wang Y.-Y., Cheng Y.-H., Chen K.-E., Tsay Y.-F. Nitrate transport, signaling, and use efficiency. Annu. Rev. Plant Biol. 2018;69:85-122.

    Not in our library10.1146/annurev-arplant-042817-040056

  35. 35.

    Kiba T., Feria-Bourrellier A.-B., Lafouge F., Lezhneva L., Boutet-Mercey S., Orsel M., Bréhaut V., Miller A., Daniel-Vedele F., Sakakibara H. The Arabidopsis nitrate transporter NRT2.4 plays a double role in roots and shoots of nitrogen-starved plants. Plant Cell. 2012;24:245-258.

    Not in our library10.1105/tpc.111.092221

  36. 36.

    Chattha M.S., Ali Q., Haroon M., Afzal M.J., Javed T., Hussain S., Mahmood T., Solanki M.K., Umar A., Abbas W. Enhancement of nitrogen use efficiency through agronomic and molecular based approaches in cotton. Front. Plant Sci. 2022;13:994306.

    Not in our library10.3389/fpls.2022.994306

  37. 37.

    Bouguyon E., Brun F., Meynard D., Kubeš M., Pervent M., Leran S., Lacombe B., Krouk G., Guiderdoni E., Zažímalová E. Multiple mechanisms of nitrate sensing by Arabidopsis nitrate transceptor NRT1. 1. Nat. Plants. 2015;1:15015.

    Not in our library10.1038/nplants.2015.15

  38. 38.

    Ho C.-H., Lin S.-H., Hu H.-C., Tsay Y.-F. CHL1 functions as a nitrate sensor in plants. Cell. 2009;138:1184-1194.

    Not in our library10.1016/j.cell.2009.07.004

  39. 39.

    Hu H.C., Wang Y.Y., Tsay Y.F. AtCIPK8, a CBL-interacting protein kinase, regulates the low-affinity phase of the primary nitrate response. Plant J. 2009;57:264-278.

    Not in our library10.1111/j.1365-313x.2008.03685.x

  40. 40.

    Maghiaoui A., Gojon A., Bach L. NRT1. 1-centered nitrate signaling in plants. J. Exp. Bot. 2020;71:6226-6237.

    Not in our library10.1093/jxb/eraa361

  41. 41.

    Izmailov S., Nikitin A. Nitrate signaling in plants: Mechanisms of implementation. Russ. J. Plant Physiol. 2020;67:31-44.

    Not in our library10.1134/s1021443720010082

  42. 42.

    O’Brien J.A., Vega A., Bouguyon E., Krouk G., Gojon A., Coruzzi G., Gutiérrez R.A. Nitrate transport, sensing, and responses in plants. Mol. Plant. 2016;9:837-856.

    Not in our library10.1016/j.molp.2016.05.004

  43. 43.

    Fang X.Z., Fang S.Q., Ye Z.Q., Liu D., Zhao K.L., Jin C.W. NRT1.1 dual-affinity nitrate transport/signalling and its roles in plant abiotic stress resistance. Front. Plant Sci. 2021;12:715694.

    Not in our library10.3389/fpls.2021.715694

  44. 44.

    Chen C.-Z., Lv X.-F., Li J.-Y., Yi H.-Y., Gong J.-M. Arabidopsis NRT1.5 is another essential component in the regulation of nitrate reallocation and stress tolerance. Plant Physiol. 2012;159:1582-1590.

    Not in our library10.1104/pp.112.199257

  45. 45.

    Zhang G.-B., Meng S., Gong J.-M. The expected and unexpected roles of nitrate transporters in plant abiotic stress resistance and their regulation. Int. J. Mol. Sci. 2018;19:3535.

    Not in our library10.3390/ijms19113535

  46. 46.

    Nazish T., Arshad M., Jan S.U., Javaid A., Khan M.H., Naeem M.A., Baber M., Ali M. Transporters and transcription factors gene families involved in improving nitrogen use efficiency (NUE) and assimilation in rice (Oryza sativa L.). Transgenic Res. 2021;31:23-42.

    Not in our library10.1007/s11248-021-00284-5

  47. 47.

    Guo B., Li Y., Wang S., Li D., Lv C., Xu R. Characterization of the Nitrate Transporter gene family and functional identification of HvNRT2.1 in barley (Hordeum vulgare L.). PLoS ONE. 2020;15:e0232056.

    Not in our library10.1371/journal.pone.0232056

  48. 48.

    Aluko O.O., Kant S., Adedire O.M., Li C., Yuan G., Liu H., Wang Q. Unlocking the potentials of nitrate transporters at improving plant nitrogen use efficiency. Front. Plant Sci. 2023;14:1074839.

    Not in our library10.3389/fpls.2023.1074839

  49. 49.

    Ji L., Song L., Zou L., Li S., Zhang R., Yang J., Wang C., Zhang Y., Wang X., Yun L. Cassava Nitrate Transporter NPF5.4 Promotes Both Yield Potential and Salt Tolerance in Rice. Mol. Breed. 2022.

    Not in our library10.21203/rs.3.rs-2206489/v1

  50. 50.

    Zhao L., Chen P., Liu P., Song Y., Zhang D. Genetic effects and expression patterns of the nitrate transporter (NRT) gene family in Populus tomentosa. Front. Plant Sci. 2021;12:661635.

    Not in our library10.3389/fpls.2021.661635

  51. 51.

    Qi J., Yu L., Ding J., Ji C., Wang S., Wang C., Ding G., Shi L., Xu F., Cai H. Transcription factor OsSNAC1 positively regulates nitrate transporter gene expression in rice. Plant Physiol. 2023;192:2923-2942.

    Not in our library10.1093/plphys/kiad290

  52. 52.

    Peña P.A., Quach T., Sato S., Ge Z., Nersesian N., Changa T., Dweikat I., Soundararajan M., Clemente T.E. Expression of the maize Dof1 transcription factor in wheat and sorghum. Front. Plant Sci. 2017;8:434.

    Not in our library10.3389/fpls.2017.00434

  53. 53.

    Cao H., Qi S., Sun M., Li Z., Yang Y., Crawford N.M., Wang Y. Overexpression of the maize ZmNLP6 and ZmNLP8 can complement the Arabidopsis nitrate regulatory mutant nlp7 by restoring nitrate signaling and assimilation. Front. Plant Sci. 2017;8:1703.

    Not in our library10.3389/fpls.2017.01703

  54. 54.

    Alvarez J.M., Riveras E., Vidal E.A., Gras D.E., Contreras-López O., Tamayo K.P., Aceituno F., Gómez I., Ruffel S., Lejay L. Systems approach identifies TGA1 and TGA4 transcription factors as important regulatory components of the nitrate response of Arabidopsis thaliana roots. Plant J. 2014;80:1-13.

    Not in our library10.1111/tpj.12618

  55. 55.

    Liu M., Chang W., Fan Y., Sun W., Qu C., Zhang K., Liu L., Xu X., Tang Z., Li J. Genome-wide identification and characterization of NODULE-INCEPTION-like protein (NLP) family genes in Brassica napus. Int. J. Mol. Sci. 2018;19:2270.

    Not in our library10.3390/ijms19082270

  56. 56.

    Chen X., Yao Q., Gao X., Jiang C., Harberd N.P., Fu X. Shoot-to-root mobile transcription factor HY5 coordinates plant carbon and nitrogen acquisition. Curr. Biol. 2016;26:640-646.

    Not in our library10.1016/j.cub.2015.12.066

  57. 57.

    Rubin G., Tohge T., Matsuda F., Saito K., Scheible W.-R.d. Members of the LBD family of transcription factors repress anthocyanin synthesis and affect additional nitrogen responses in Arabidopsis. Plant Cell. 2009;21:3567-3584.

    Not in our library10.1105/tpc.109.067041

  58. 58.

    Vidal E.A., Álvarez J.M., Gutiérrez R.A. Nitrate regulation of AFB3 and NAC4 gene expression in Arabidopsis roots depends on NRT1. 1 nitrate transport function. Plant Signal. Behav. 2014;9:e28501.

    Not in our library10.4161/psb.28501

  59. 59.

    Du X.-Q., Wang F.-L., Li H., Jing S., Yu M., Li J., Wu W.-H., Kudla J., Wang Y. The transcription factor MYB59 regulates K+/NO3− translocation in the Arabidopsis response to low K+ stress. Plant Cell. 2019;31:699-714.

    Not in our library10.1105/tpc.18.00674

  60. 60.

    Medici A., Marshall-Colon A., Ronzier E., Szponarski W., Wang R., Gojon A., Crawford N.M., Ruffel S., Coruzzi G.M., Krouk G. AtNIGT1/HRS1 integrates nitrate and phosphate signals at the Arabidopsis root tip. Nat. Commun. 2015;6:6274.

    Not in our library10.1038/ncomms7274

  61. 61.

    Guan P., Wang R., Nacry P., Breton G., Kay S.A., Pruneda-Paz J.L., Davani A., Crawford N.M. Nitrate foraging by Arabidopsis roots is mediated by the transcription factor TCP20 through the systemic signaling pathway. Proc. Natl. Acad. Sci. USA. 2014;111:15267-15272.

    Not in our library10.1073/pnas.1411375111

  62. 62.

    Dash M., Yordanov Y.S., Georgieva T., Kumari S., Wei H., Busov V. A systems biology approach identifies new regulators of poplar root development under low nitrogen. Plant J. 2015;84:335-346.

    Not in our library10.1111/tpj.13002

  63. 63.

    Shin J.M., Chung K., Sakamoto S., Kojima S., Yeh C.-M., Ikeda M., Mitsuda N., Ohme-Takagi M. The chimeric repressor for the GATA4 transcription factor improves tolerance to nitrogen deficiency in Arabidopsis. Plant Biotechnol. 2017;34:151-158.

    Not in our library10.5511/plantbiotechnology.17.0727a

  64. 64.

    Chen M., Zhu K., Xie J., Liu J., Tan P., Peng F. Genome-Wide Identification and Expression Analysis of AMT and NRT Gene Family in Pecan (Carya illinoinensis) Seedlings Revealed a Preference for NH4+-N. Int. J. Mol. Sci. 2022;23:13314.

    Not in our library10.3390/ijms232113314

  65. 65.

    Schuldiner S., Shirvan A., Linial M. Vesicular neurotransmitter transporters: From bacteria to humans. Physiol. Rev. 1995;75:369-392.

    Not in our library10.1152/physrev.1995.75.2.369

  66. 66.

    Tsay Y.-F., Chiu C.-C., Tsai C.-B., Ho C.-H., Hsu P.-K. Nitrate transporters and peptide transporters. FEBS Lett. 2007;581:2290-2300.

    Not in our library10.1016/j.febslet.2007.04.047

  67. 67.

    Sun J., Bankston J.R., Payandeh J., Hinds T.R., Zagotta W.N., Zheng N. Crystal structure of the plant dual-affinity nitrate transporter NRT1. 1. Nature. 2014;507:73-77.

    Not in our library10.1038/nature13074

  68. 68.

    Madej M.G., Dang S., Yan N., Kaback H.R. Evolutionary mix-and-match with MFS transporters. Proc. Natl. Acad. Sci. USA. 2013;110:5870-5874.

    Not in our library10.1073/pnas.1303538110

  69. 69.

    Orsel M., Chopin F., Leleu O., Smith S.J., Krapp A., Daniel-Vedele F.o., Miller A.J. Characterization of a Two-Component High-Affinity Nitrate Uptake System in Arabidopsis. Physiology and Protein-Protein Interaction. Plant Physiol. 2006;142:1304-1317.

    Not in our library10.1104/pp.106.085209

  70. 70.

    Okamoto M., Kumar A., Li W., Wang Y., Siddiqi M.Y., Crawford N.M., Glass A.D.M. High-Affinity Nitrate Transport in Roots of Arabidopsis Depends on Expression of the NAR2-Like Gene AtNRT3.1. Plant Physiol. 2006;140:1036-1046.

    Not in our library10.1104/pp.105.074385

  71. 71.

    Benková E., Bielach A. Lateral root organogenesis—From cell to organ. Curr. Opin. Plant Biol. 2010;13:677-683.

    Not in our library10.1016/j.pbi.2010.09.006

  72. 72.

    Yong Z., Kotur Z., Glass A.D. Characterization of an intact two-component high-affinity nitrate transporter from Arabidopsis roots. Plant J. 2010;63:739-748.

    Not in our library10.1111/j.1365-313x.2010.04278.x

  73. 73.

    Bowen G.D., Smith S.E. The effects of mycorrhizas on nitrogen uptake by plants. Ecol. Bull. 1981;33:237-247.

    Not in our library

  74. 74.

    Öhlund J., Näsholm T. Growth of conifer seedlings on organic and inorganic nitrogen sources. Tree Physiol. 2001;21:1319-1326.

    Not in our library10.1093/treephys/21.18.1319

  75. 75.

    Gazzarrini S., Lejay L., Gojon A., Ninnemann O., Frommer W.B., von Wirén N. Three Functional Transporters for Constitutive, Diurnally Regulated, and Starvation-Induced Uptake of Ammonium into Arabidopsis Roots. Plant Cell. 1999;11:937-947.

    Not in our library10.1105/tpc.11.5.937

  76. 76.

    Bloom A.J., Meyerhoff P.A., Taylor A.R., Rost T.L. Root Development and Absorption of Ammonium and Nitrate from the Rhizosphere. J. Plant Growth Regul. 2002;21:416-431.

    Not in our library10.1007/s00344-003-0009-8

  77. 77.

    Bittsánszky A., Pilinszky K., Gyulai G., Komives T. Overcoming ammonium toxicity. Plant Sci. 2015;231:184-190.

    Not in our library10.1016/j.plantsci.2014.12.005

  78. 78.

    Renard J.J., Calidonna S.E., Henley M.V. Fate of ammonia in the atmosphere—A review for applicability to hazardous releases. J. Hazard. Mater. 2004;108:29-60.

    Not in our library10.1016/j.jhazmat.2004.01.015

  79. 79.

    Hachiya T., Watanabe C.K., Fujimoto M., Ishikawa T., Takahara K., Kawai-Yamada M., Uchimiya H., Uesono Y., Terashima I., Noguchi K. Nitrate Addition Alleviates Ammonium Toxicity Without Lessening Ammonium Accumulation, Organic Acid Depletion and Inorganic Cation Depletion in Arabidopsis thaliana Shoots. Plant Cell Physiol. 2012;53:577-591.

    Not in our library10.1093/pcp/pcs012

  80. 80.

    von Wirén N., Gazzarrini S., Gojon A., Frommer W.B. The molecular physiology of ammonium uptake and retrieval. Curr. Opin. Plant Biol. 2000;3:254-261.

    Not in our library10.1016/s1369-5266(00)00073-x

  81. 81.

    Loqué D., von Wirén N. Regulatory levels for the transport of ammonium in plant roots. J. Exp. Bot. 2004;55:1293-1305.

    Not in our library10.1093/jxb/erh147

  82. 82.

    Glass A.D.M., Britto D.T., Kaiser B.N., Kinghorn J.R., Kronzucker H.J., Kumar A., Okamoto M., Rawat S., Siddiqi M.Y., Unkles S.E., et al. The regulation of nitrate and ammonium transport systems in plants. J. Exp. Bot. 2002;53:855-864.

    Not in our library10.1093/jexbot/53.370.855

  83. 83.

    Hao D.-L., Zhou J.-Y., Yang S.-Y., Qi W., Yang K.-J., Su Y.-H. Function and Regulation of Ammonium Transporters in Plants. Int. J. Mol. Sci. 2020;21:3557.

    Not in our library10.3390/ijms21103557

  84. 84.

    Behie S.W., Bidochka M.J. Ubiquity of Insect-Derived Nitrogen Transfer to Plants by Endophytic Insect-Pathogenic Fungi: An Additional Branch of the Soil Nitrogen Cycle. Appl. Environ. Microbiol. 2014;80:1553-1560.

    Not in our library10.1128/aem.03338-13

  85. 85.

    Giehl R.F.H., Laginha A.M., Duan F., Rentsch D., Yuan L., von Wirén N. A Critical Role of AMT2;1 in Root-To-Shoot Translocation of Ammonium in Arabidopsis. Mol. Plant. 2017;10:1449-1460.

    Not in our library10.1016/j.molp.2017.10.001

  86. 86.

    Couturier J., Montanini B., Martin F., Brun A., Blaudez D., Chalot M. The expanded family of ammonium transporters in the perennial poplar plant. New Phytol. 2007;174:137-150.

    Not in our library10.1111/j.1469-8137.2007.01992.x

  87. 87.

    Esmaeilzadeh-Salestani K., Samandari_Bahraseman M.R., Tohidfar M., Khaleghdoust B., Keres I., Mõttus A., Loit E. Expression of AMT1;1 and AMT2;1 is stimulated by mineral nitrogen and reproductive growth stage in barley under field conditions. J. Plant Nutr. 2023;46:1246-1258.

    Not in our library10.1080/01904167.2022.2067764

  88. 88.

    Wu X., Yang H., Qu C., Xu Z., Li W., Hao B., Yang C., Sun G., Liu G. Sequence and expression analysis of the AMT gene family in poplar. Front. Plant Sci. 2015;6:337.

    Not in our library10.3389/fpls.2015.00337

  89. 89.

    Wu X.X., Yuan D.P., Chen H., Kumar V., Kang S.M., Jia B., Xuan Y.H. Ammonium transporter 1 increases rice resistance to sheath blight by promoting nitrogen assimilation and ethylene signalling. Plant Biotechnol. J. 2022;20:1085-1097.

    Not in our library10.1111/pbi.13789

  90. 90.

    Pastor V., Gamir J., Camañes G., Cerezo M., Sánchez-Bel P., Flors V. Disruption of the ammonium transporter AMT1.1 alters basal defenses generating resistance against Pseudomonas syringae and Plectosphaerella cucumerina. Front. Plant Sci. 2014;5:231.

    Not in our library10.3389/fpls.2014.00231

  91. 91.

    Lima J.E., Kojima S., Takahashi H., von Wirén N. Ammonium Triggers Lateral Root Branching in Arabidopsis in an AMMONIUM TRANSPORTER1;3-Dependent Manner. Plant Cell. 2010;22:3621-3633.

    Not in our library10.1105/tpc.110.076216

  92. 92.

    Rogato A., D’Apuzzo E., Chiurazzi M. The multiple plant response to high ammonium conditions. Plant Signal. Behav. 2010;5:1594-1596.

    Not in our library10.4161/psb.5.12.13856

  93. 93.

    Rogato A., D’Apuzzo E., Barbulova A., Omrane S., Parlati A., Carfagna S., Costa A., Schiavo F.L., Esposito S., Chiurazzi M. Characterization of a Developmental Root Response Caused by External Ammonium Supply in Lotus japonicus. Plant Physiol. 2010;154:784-795.

    Not in our library10.1104/pp.110.160309

  94. 94.

    Cao H., Liu Q., Liu X., Ma Z., Zhang J., Li X., Shen L., Yuan J., Zhang Q. Phosphatidic acid regulates ammonium uptake by interacting with AMMONIUM TRANSPORTER 1;1 in Arabidopsis. Plant Physiol. 2023.

    Not in our library10.1093/plphys/kiad421

  95. 95.

    Giovannetti M., Volpe V., Salvioli A., Bonfante P., Johnson N.C., Gehring C., Jansa J. Chapter 7—Fungal and Plant Tools for the Uptake of Nutrients in Arbuscular Mycorrhizas: A Molecular View. Mycorrhizal Mediation of Soil. 2017;107-128.

    Not in our library

  96. 96.

    Wu Y., Yang W., Wei J., Yoon H., An G. Transcription Factor OsDOF18 Controls Ammonium Uptake by Inducing Ammonium Transporters in Rice Roots. Mol. Cells. 2017;40:178-185.

    Not in our library10.14348/molcells.2017.2261

  97. 97.

    Yuan L., Gu R., Xuan Y., Smith-Valle E., Loqué D., Frommer W.B., von Wirén N. Allosteric Regulation of Transport Activity by Heterotrimerization of Arabidopsis Ammonium Transporter Complexes in Vivo. Plant Cell. 2013;25:974-984.

    Not in our library10.1105/tpc.112.108027

  98. 98.

    Lanquar V., Loqué D., Hörmann F., Yuan L., Bohner A., Engelsberger W.R., Lalonde S., Schulze W.X., von Wireén N., Frommer W.B. Feedback Inhibition of Ammonium Uptake by a Phospho-Dependent Allosteric Mechanism in Arabidopsis. Plant Cell. 2009;21:3610-3622.

    Not in our library10.1105/tpc.109.068593

  99. 99.

    Loqué D., Lalonde S., Looger L.L., von Wirén N., Frommer W.B. A cytosolic trans-activation domain essential for ammonium uptake. Nature. 2007;446:195-198.

    Not in our library10.1038/nature05579

  100. 100.

    Søgaard R., Alsterfjord M., MacAulay N., Zeuthen T. Ammonium ion transport by the AMT/Rh homolog TaAMT1;1 is stimulated by acidic pH. Pflügers Arch.—Eur. J. Physiol. 2009;458:733-743.

    Not in our library10.1007/s00424-009-0665-z

  101. 101.

    Wang Q., Zhao Y., Luo W., Li R., He Q., Fang X., Michele R.D., Ast C., von Wirén N., Lin J. Single-particle analysis reveals shutoff control of the Arabidopsis ammonium transporter AMT1;3 by clustering and internalization. Proc. Natl. Acad. Sci. USA. 2013;110:13204-13209.

    Not in our library10.1073/pnas.1301160110

  102. 102.

    Koltun A., Maniero R.A., Vitti M., de Setta N., Giehl R.F.H., Lima J.E., Figueira A. Functional characterization of the sugarcane (Saccharum spp.) ammonium transporter AMT2;1 suggests a role in ammonium root-to-shoot translocation. Front. Plant Sci. 2022;13:1039041.

    Not in our library10.3389/fpls.2022.1039041

  103. 103.

    Clegg S., Yu F., Griffiths L., Cole J.A. The roles of the polytopic membrane proteins NarK, NarU and NirC in Escherichia coli K-12: Two nitrate and three nitrite transporters. Mol. Microbiol. 2002;44:143-155.

    Not in our library10.1046/j.1365-2958.2002.02858.x

  104. 104.

    Wang Y., Li W., Siddiqi Y., Symington V.F., Kinghorn J.R., Unkles S.E., Glass A.D. Nitrite transport is mediated by the nitrite-specific high-affinity NitA transporter and by nitrate transporters NrtA, NrtB in Aspergillus nidulans. Fungal Genet. Biol. 2008;45:94-102.

    Not in our library10.1016/j.fgb.2007.10.001

  105. 105.

    Kotur Z., Siddiqi Y.M., Glass A.D. Characterization of nitrite uptake in Arabidopsis thaliana: Evidence for a nitrite-specific transporter. New Phytol. 2013;200:201-210.

    Not in our library10.1111/nph.12358

  106. 106.

    Sugiura M., Georgescu M.N., Takahashi M. A nitrite transporter associated with nitrite uptake by higher plant chloroplasts. Plant Cell Physiol. 2007;48:1022-1035.

    Not in our library10.1093/pcp/pcm073

  107. 107.

    Maeda S.-I., Konishi M., Yanagisawa S., Omata T. Nitrite transport activity of a novel HPP family protein conserved in cyanobacteria and chloroplasts. Plant Cell Physiol. 2014;55:1311-1324.

    Not in our library10.1093/pcp/pcu075

  108. 108.
    Pfam: The protein families database in 2021 · 2020
    No notice
  109. 109.

    Mengel K., Kirkby E.A., Kosegarten H., Appel T. Nitrogen. Principles of Plant Nutrition. 2001;397-434.

    Not in our library

  110. 110.

    Sparacino-Watkins C., Stolz J.F., Basu P. Nitrate and periplasmic nitrate reductases. Chem. Soc. Rev. 2014;43:676-706.

    Not in our library10.1039/c3cs60249d

  111. 111.

    Rosswall T., Likens G.E. Some Perspectives of the Major Biogeochemical Cycles. 1981;Volume 2.

    Not in our library

  112. 112.
    Workgroup Report: Drinking-Water Nitrate and Health—Recent Findings and Research Needs · 2005
    No notice
  113. 113.

    Powlson D.S., Addiscott T.M., Benjamin N., Cassman K.G., de Kok T.M., van Grinsven H., L’hirondel J.L., Avery A.A., Van Kessel C. When does nitrate become a risk for humans? J. Environ. Qual. 2008;37:291-295.

    Not in our library10.2134/jeq2007.0177

  114. 114.

    Bonner F.T., Hughes M.N. The aqueous solution chemistry of nitrogen in low positive oxidation states. Comments Inorg. Chem. 1988;7:215-234.

    Not in our library10.1080/02603598808072309

  115. 115.

    Jaffe D.A., Butcher S.S., Charlson R.J., Orians G.H., Wolfe G.V. 12 The Nitrogen Cycle. International Geophysics. 1992;Volume 50:263-284.

    Not in our library

  116. 116.

    Canfield D.E., Glazer A.N., Falkowski P.G. The evolution and future of Earth’s nitrogen cycle. Science. 2010;330:192-196.

    Not in our library10.1126/science.1186120

  117. 117.

    Ishii S., Ikeda S., Minamisawa K., Senoo K. Nitrogen cycling in rice paddy environments: Past achievements and future challenges. Microbes Environ. 2011;26:282-292.

    Not in our library10.1264/jsme2.me11293

  118. 118.

    Elrys A.S., Zhu Q., Jiang C., Liu J., Sobhy H.H., Shen Q., Uwiragiye Y., Wu Y., El-Tarabily K.A., Meng L. Global soil nitrogen cycle pattern and nitrogen enrichment effects: Tropical versus subtropical forests. Glob. Chang. Biol. 2023;29:1905-1921.

    Not in our library10.1111/gcb.16603

  119. 119.

    Joerger R., Jacobson M., Premakumar R., Wolfinger E., Bishop P. Nucleotide sequence and mutational analysis of the structural genes (anfHDGK) for the second alternative nitrogenase from Azotobacter vinelandii. J. Bacteriol. 1989;171:1075-1086.

    Not in our library10.1128/jb.171.2.1075-1086.1989

  120. 120.

    Joerger R., Loveless T., Pau R., Mitchenall L., Simon B., Bishop P. Nucleotide sequences and mutational analysis of the structural genes for nitrogenase 2 of Azotobacter vinelandii. J. Bacteriol. 1990;172:3400-3408.

    Not in our library10.1128/jb.172.6.3400-3408.1990

  121. 121.

    Fani R., Gallo R., Liò P. Molecular evolution of nitrogen fixation: The evolutionary history of the nifD, nifK, nifE, and nifN genes. J. Mol. Evol. 2000;51:1-11.

    Not in our library10.1007/s002390010061

  122. 122.

    Cantera J.J.L., Stein L.Y. Molecular diversity of nitrite reductase genes (nirK) in nitrifying bacteria. Environ. Microbiol. 2007;9:765-776.

    Not in our library10.1111/j.1462-2920.2006.01198.x

  123. 123.

    Kartal B., Maalcke W.J., De Almeida N.M., Cirpus I., Gloerich J., Geerts W., Op den Camp H.J., Harhangi H.R., Janssen-Megens E.M., Francoijs K.-J. Molecular mechanism of anaerobic ammonium oxidation. Nature. 2011;479:127-130.

    Not in our library10.1038/nature10453

  124. 124.

    Jepson B.J., Mohan S., Clarke T.A., Gates A.J., Cole J.A., Butler C.S., Butt J.N., Hemmings A.M., Richardson D.J. Spectropotentiometric and structural analysis of the periplasmic nitrate reductase from Escherichia coli. J. Biol. Chem. 2007;282:6425-6437.

    Not in our library10.1074/jbc.m607353200

  125. 125.

    Heiss B., Frunzke K., Zumft W. Formation of the NN bond from nitric oxide by a membrane-bound cytochrome bc complex of nitrate-respiring (denitrifying) Pseudomonas stutzeri. J. Bacteriol. 1989;171:3288-3297.

    Not in our library10.1128/jb.171.6.3288-3297.1989

  126. 126.

    Suzuki M., Arai H., Ishii M., Igarashi Y. Gene structure and expression profile of cytochrome bc nitric oxide reductase from Hydrogenobacter thermophilus TK-6. Biosci. Biotechnol. Biochem. 2006;70:1666-1671.

    Not in our library10.1271/bbb.60018

  127. 127.

    Ogawa K.-i., Akagawa E., Yamane K., Sun Z.-W., LaCelle M., Zuber P., Nakano M.M. The nasB operon and nasA gene are required for nitrate/nitrite assimilation in Bacillus subtilis. J. Bacteriol. 1995;177:1409-1413.

    Not in our library10.1128/jb.177.5.1409-1413.1995

  128. 128.

    Exley G.E., Colandene J.D., Garrett R.H. Molecular cloning, characterization, and nucleotide sequence of nit-6, the structural gene for nitrite reductase in Neurospora crassa. J. Bacteriol. 1993;175:2379-2392.

    Not in our library10.1128/jb.175.8.2379-2392.1993

  129. 129.

    Sohaskey C.D., Wayne L.G. Role of narK2X and narGHJI inHypoxic Upregulation of Nitrate Reduction by Mycobacteriumtuberculosis. J. Bacteriol. 2003;185:7247-7256.

    Not in our library10.1128/jb.185.24.7247-7256.2003

  130. 130.

    Stolz J.F., Basu P. Evolution of nitrate reductase: Molecular and structural variations on a common function. Chembiochem. 2002;3:198-206.

    Not in our library10.1002/1439-7633(20020301)3:2/3<198::aid-cbic198>3.0.co;2-c

  131. 131.

    Kizawa H., Tomura D., Oda M., Fukamizu A., Hoshino T., Gotoh O., Yasui T., Shoun H. Nucleotide sequence of the unique nitrate/nitrite-inducible cytochrome P-450 cDNA from Fusarium oxysporum. J. Biol. Chem. 1991;266:10632-10637.

    Not in our library10.1016/s0021-9258(18)99270-7

  132. 132.

    Zhang L., Shoun H. Purification and functional analysis of fungal nitric oxide reductase cytochrome P450nor. Methods Enzymol. 2008;437:117-133.

    Not in our library10.1016/s0076-6879(07)37007-9

  133. 133.

    COYLE C.L., ZUMFT W.G., KRONECK P.M., KÖRNER H., JAKOB W. Nitrous oxide reductase from denitrifying: Pseudomonas perfectomarina purification and properties of a novel multicopper enzyme. Eur. J. Biochem. 1985;153:459-467.

    Not in our library10.1111/j.1432-1033.1985.tb09324.x

  134. 134.

    Arai H., Mizutani M., Igarashi Y. Transcriptional regulation of the nos genes for nitrous oxide reductase in Pseudomonas aeruginosa. Microbiology. 2003;149:29-36.

    Not in our library10.1099/mic.0.25936-0

  135. 135.

    Zumft W.G., Kroneck P.M. Respiratory transformation of nitrous oxide (N2O) to dinitrogen by Bacteria and Archaea. Adv. Microb. Physiol. 2006;52:107-227.

    Not in our library10.1016/s0065-2911(06)52003-x

  136. 136.

    Keable S.M., Vertemara J., Zadvornyy O.A., Eilers B.J., Danyal K., Rasmussen A.J., De Gioia L., Zampella G., Seefeldt L.C., Peters J.W. Structural characterization of the nitrogenase molybdenum-iron protein with the substrate acetylene trapped near the active site. J. Inorg. Biochem. 2018;180:129-134.

    Not in our library10.1016/j.jinorgbio.2017.12.008

  137. 137.

    Hu Y., Lee C.C., Ribbe M.W. Extending the carbon chain: Hydrocarbon formation catalyzed by vanadium/molybdenum nitrogenases. Science. 2011;333:753-755.

    Not in our library10.1126/science.1206883

  138. 138.

    Deistung J., Cannon F.C., Cannon M.C., Hill S., Thorneley R. Electron transfer to nitrogenase in Klebsiella pneumoniae. nifF gene cloned and the gene product, a flavodoxin, purified. Biochem. J. 1985;231:743.

    Not in our library10.1042/bj2310743

  139. 139.

    Fraser J.A., Davis M.A., Hynes M.J. The formamidase gene of Aspergillus nidulans: Regulation by nitrogen metabolite repression and transcriptional interference by an overlapping upstream gene. Genetics. 2001;157:119-131.

    Not in our library10.1093/genetics/157.1.119

  140. 140.

    Maalcke W.J., Reimann J., De Vries S., Butt J.N., Dietl A., Kip N., Mersdorf U., Barends T.R., Jetten M.S., Keltjens J.T. Characterization of anammox hydrazine dehydrogenase, a key N2-producing enzyme in the global nitrogen cycle. J. Biol. Chem. 2016;291:17077-17092.

    Not in our library10.1074/jbc.m116.735530

  141. 141.

    Zhao R., Biddle J.F., Jørgensen S.L. Introducing Candidatus Bathyanammoxibiaceae, a family of bacteria with the anammox potential present in both marine and terrestrial environments. ISME Commun. 2022;2:42.

    Not in our library10.1038/s43705-022-00125-4

  142. 142.

    Breton S. The cellular physiology of carbonic anhydrases. Jop. 2001;2:159-164.

    Not in our library

  143. 143.

    Pastorekova S., Parkkila S., Pastorek J., Supuran C. Carbonic anhydrases: Current state of the art, therapeutic applications and future prospects. J. Enzym. Inhib. Med. Chem. 2004;19:199-229.

    Not in our library10.1080/14756360410001689540

  144. 144.

    Harano Y., Suzuki I., Maeda S.-i., Kaneko T., Tabata S., Omata T. Identification and nitrogen regulation of the cyanase gene from the cyanobacteria Synechocystis sp. strain PCC 6803 and Synechococcus sp. strain PCC 7942. J. Bacteriol. 1997;179:5744-5750.

    Not in our library10.1128/jb.179.18.5744-5750.1997

  145. 145.

    Cantera J.J.L., Stein L.Y. Role of nitrite reductase in the ammonia-oxidizing pathway of Nitrosomonas europaea. Arch. Microbiol. 2007;188:349-354.

    Not in our library10.1007/s00203-007-0255-4

  146. 146.

    Noji S., Nohno T., Saito T., Taniguchi S. The narK gene product participates in nitrate transport induced in Escherichia coli nitrate-respiring cells. FEBS Lett. 1989;252:139-143.

    Not in our library10.1016/0014-5793(89)80906-8

  147. 147.

    Maeda S.-I., Omata T. Nitrite transport activity of the ABC-type cyanate transporter of the cyanobacterium Synechococcus elongatus. J. Bacteriol. 2009;191:3265-3272.

    Not in our library10.1128/jb.00013-09

  148. 148.

    Blasco F., Iobbi C., Ratouchniak J., Bonnefoy V., Chippaux M. Nitrate reductases of Escherichia coli: Sequence of the second nitrate reductase and comparison with that encoded by the narGHJI operon. Mol. Gen. Genet. MGG. 1990;222:104-111.

    Not in our library10.1007/bf00283030

  149. 149.

    Okamoto P.M., Marzluf G.A. Nitrate reductase of Neurospora crassa: The functional role of individual amino acids in the heme domain as examined by site-directed mutagenesis. Mol. Gen. Genet. MGG. 1993;240:221-230.

    Not in our library10.1007/bf00277060

  150. 150.

    Rubio L.M., Herrero A., Flores E. A cyanobacterial narB gene encodes a ferredoxin-dependent nitrate reductase. Plant Mol. Biol. 1996;30:845-850.

    Not in our library10.1007/bf00019017

  151. 151.

    Takahashi M., Sasaki Y., Ida S., Morikawa H. Nitrite reductase gene enrichment improves assimilation of NO2 in Arabidopsis. Plant Physiol. 2001;126:731-741.

    Not in our library10.1104/pp.126.2.731

  152. 152.

    Harborne N.R., Griffiths L., Busby S.J., Cole J.A. Transcriptional control, translation and function of the products of the five open reading frames of the Escherichia coli nir operon. Mol. Microbiol. 1992;6:2805-2813.

    Not in our library10.1111/j.1365-2958.1992.tb01460.x

  153. 153.

    Einsle O., Messerschmidt A., Stach P., Bourenkov G.P., Bartunik H.D., Huber R., Kroneck P.M. Structure of cytochrome c nitrite reductase. Nature. 1999;400:476-480.

    Not in our library10.1038/22802

  154. 154.

    Gadda G., Francis K. Nitronate monooxygenase, a model for anionic flavin semiquinone intermediates in oxidative catalysis. Arch. Biochem. Biophys. 2010;493:53-61.

    Not in our library10.1016/j.abb.2009.06.018

  155. 155.

    Daubner S.C., Gadda G., Valley M.P., Fitzpatrick P.F. Cloning of nitroalkane oxidase from Fusarium oxysporum identifies a new member of the acyl-CoA dehydrogenase superfamily. Proc. Natl. Acad. Sci. USA. 2002;99:2702-2707.

    Not in our library10.1073/pnas.052527799

  156. 156.

    Wolfe M.T., Heo J., Garavelli J.S., Ludden P.W. Hydroxylamine Reductase Activity of the Hybrid Cluster Protein from Escherichia coli. J. Bacteriol. 2002;184:5898-5902.

    Not in our library10.1128/jb.184.21.5898-5902.2002

  157. 157.

    Hommes N.G., Sayavedra-Soto L.A., Arp D.J. Transcript Analysis of Multiple Copies of amo (Encoding Ammonia Monooxygenase) and hao (Encoding Hydroxylamine Oxidoreductase) in Nitrosomonas europaea. J. Bacteriol. 2001;183:1096-1100.

    Not in our library10.1128/jb.183.3.1096-1100.2001

  158. 158.

    MOIR J.W.B., WEHRFRITZ J.-M., SPIRO S., RICHARDSON D.J. The biochemical characterization of a novel non-haem-iron hydroxylamine oxidase from Paracoccus denitrificans GB17. Biochem. J. 1996;319:823-827.

    Not in our library10.1042/bj3190823

  159. 159.

    Klotz M.G., Stein L.Y. Genomics of Ammonia-Oxidizing Bacteria and Insights into Their Evolution. Nitrification. 2011;57-94.

    Not in our library10.1128/9781555817145.ch4

  160. 160.

    Durbecq V., Legrain C., Roovers M., Piérard A., Glansdorff N. The carbamate kinase-like carbamoyl phosphate synthetase of the hyperthermophilic archaeon Pyrococcus furiosus, a missing link in the evolution of carbamoyl phosphate biosynthesis. Proc. Natl. Acad. Sci. USA. 1997;94:12803-12808.

    Not in our library10.1073/pnas.94.24.12803

  161. 161.

    Ahuja V., Powers-Lee S.G. Human carbamoyl-phosphate synthetase: Insight into N-acetylglutamate interaction and the functional effects of a common single nucleotide polymorphism. J. Inherit. Metab. Dis. 2008;31:481-491.

    Not in our library10.1007/s10545-008-0913-y

  162. 162.

    Belitsky B.R., Sonenshein A.L. Role and Regulation of Bacillus subtilis Glutamate Dehydrogenase Genes. J. Bacteriol. 1998;180:6298-6305.

    Not in our library10.1128/jb.180.23.6298-6305.1998

  163. 163.

    Frigerio F., Casimir M., Carobbio S., Maechler P. Tissue specificity of mitochondrial glutamate pathways and the control of metabolic homeostasis. Biochim. Biophys. Acta (BBA)-Bioenerg. 2008;1777:965-972.

    Not in our library10.1016/j.bbabio.2008.04.031

  164. 164.

    Riba L., Becerril B., Servín-González L., Valle F., Bolivar F. Identification of a functional promoter for the Escherichia coli gdhA gene and its regulation. Gene. 1988;71:233-246.

    Not in our library10.1016/0378-1119(88)90040-6

  165. 165.

    Newsholme P., Procopio J., Lima M.M.R., Pithon-Curi T.C., Curi R. Glutamine and glutamate—their central role in cell metabolism and function. Cell Biochem. Funct. 2003;21:1-9.

    Not in our library10.1002/cbf.1003

  166. 166.

    Zhu D., Mukherjee C., Yang Y., Rios B.E., Gallagher D.T., Smith N.N., Biehl E.R., Hua L. A new nitrilase from Bradyrhizobium japonicum USDA 110: Gene cloning, biochemical characterization and substrate specificity. J. Biotechnol. 2008;133:327-333.

    Not in our library10.1016/j.jbiotec.2007.10.001

  167. 167.

    Sonawane A. Regulation of Pseudomonas putida Genes Involved in the Metabolism of Acidic Amino Acids. Ph.D. Thesis. 2003.

    Not in our library

  168. 168.

    Lancien M., Martin M., Hsieh M.-H., Leustek T., Goodman H., Coruzzi G.M. Arabidopsis glt1-T mutant defines a role for NADH-GOGAT in the non-photorespiratory ammonium assimilatory pathway. Plant J. 2002;29:347-358.

    Not in our library10.1046/j.1365-313x.2002.01218.x

  169. 169.

    Coschigano K.T., Melo-Oliveira R., Lim J., Coruzzi G.M. Arabidopsis gls Mutants and Distinct Fd-GOGAT Genes: Implications for Photorespiration and Primary Nitrogen Assimilation. Plant Cell. 1998;10:741-752.

    Not in our library10.1105/tpc.10.5.741

  170. 170.

    Rees E., Siddiqui R.A., Köster F., Schneider B., Friedrich B. Structural gene (nirS) for the cytochrome cd1 nitrite reductase of Alcaligenes eutrophus H16. Appl. Environ. Microbiol. 1997;63:800-802.

    Not in our library10.1128/aem.63.2.800-802.1997

  171. 171.

    Hole U.H., Vollack K.-U., Zumft W.G., Eisenmann E., Siddiqui R.A., Friedrich B., Kroneck P.M.H. Characterization of the membranous denitrification enzymes nitrite reductase (cytochrome cd1) and copper-containing nitrous oxide reductase from Thiobacillus denitrificans. Arch. Microbiol. 1996;165:55-61.

    Not in our library10.1007/s002030050296

  172. 172.

    Dubey R.S., Srivastava R.K., Pessarakli M. Physiological mechanisms of nitrogen absorption and assimilation in plants under stressful conditions. Handbook of Plant and Crop Physiology. 2021;579-616.

    Not in our library

  173. 173.

    Straub T., Ludewig U., Neuhäuser B. The kinase CIPK23 inhibits ammonium transport in Arabidopsis thaliana. Plant Cell. 2017;29:409-422.

    Not in our library10.1105/tpc.16.00806

  174. 174.

    Engels C., Marschner H. Plant uptake and utilization of nitrogen. Nitrogen Fertilization in the Environment. 1995;41-81.

    Not in our library

  175. 175.

    Garg S. Role and hormonal regulation of nitrate reductase activity in higher plants: A review. Plant Sci. Feed. 2013;3:13-20.

    Not in our library

  176. 176.

    Kaiser W.M., Huber S.C. Post-translational regulation of nitrate reductase: Mechanism, physiological relevance and environmental triggers. J. Exp. Bot. 2001;52:1981-1989.

    Not in our library10.1093/jexbot/52.363.1981

  177. 177.

    Nemie-Feyissa D., Królicka A., Førland N., Hansen M., Heidari B., Lillo C. Post-translational control of nitrate reductase activity responding to light and photosynthesis evolved already in the early vascular plants. J. Plant Physiol. 2013;170:662-667.

    Not in our library10.1016/j.jplph.2012.12.010

  178. 178.

    Balotf S., Kavoosi G., Kholdebarin B. Nitrate reductase, nitrite reductase, glutamine synthetase, and glutamate synthase expression and activity in response to different nitrogen sources in nitrogen-starved wheat seedlings. Biotechnol. Appl. Biochem. 2016;63:220-229.

    Not in our library10.1002/bab.1362

  179. 179.

    Vega-Mas I., Cukier C., Coleto I., González-Murua C., Limami A.M., González-Moro M.B., Marino D. Isotopic labelling reveals the efficient adaptation of wheat root TCA cycle flux modes to match carbon demand under ammonium nutrition. Sci. Rep. 2019;9:8925.

    Not in our library10.1038/s41598-019-45393-8

  180. 180.

    Fujita T., Beier M.P., Tabuchi-Kobayashi M., Hayatsu Y., Nakamura H., Umetsu-Ohashi T., Sasaki K., Ishiyama K., Murozuka E., Kojima M. Cytosolic glutamine synthetase GS1; 3 is involved in rice grain ripening and germination. Front. Plant Sci. 2022;13:835835.

    Not in our library10.3389/fpls.2022.835835

  181. 181.

    Lea P.J., Miflin B.J. Nitrogen assimilation and its relevance to crop improvement. Annu. Plant Rev. 2011;42:1-40.

    Not in our library

  182. 182.

    Miflin B.J., Lea P.J. The pathway of nitrogen assimilation in plants. Phytochemistry. 1976;15:873-885.

    Not in our library10.1016/s0031-9422(00)84362-9

  183. 183.

    Oaks A., Stulen I., Jones K., Winspear M.J., Misra S., Boesel I.L. Enzymes of nitrogen assimilation in maize roots. Planta. 1980;148:477-484.

    Not in our library10.1007/bf02395318

  184. 184.

    Tamura W., Hidaka Y., Tabuchi M., Kojima S., Hayakawa T., Sato T., Obara M., Kojima M., Sakakibara H., Yamaya T. Reverse genetics approach to characterize a function of NADH-glutamate synthase1 in rice plants. Amino Acids. 2010;39:1003-1012.

    Not in our library10.1007/s00726-010-0531-5

  185. 185.

    Kojima S., Minagawa H., Yoshida C., Inoue E., Takahashi H., Ishiyama K. Coregulation of glutamine synthetase1; 2 (GLN1; 2) and NADH-dependent glutamate synthase (GLT1) gene expression in Arabidopsis roots in response to ammonium supply. Front. Plant Sci. 2023;14:1127006.

    Not in our library10.3389/fpls.2023.1127006

  186. 186.

    Yasuda T., Konishi N., Kojima S. Ammonium uptake capacity and response of cytosolic glutamine synthetase 1;2 to ammonium supply are key factors for the adaptation of ammonium nutrition in Arabidopsis thaliana. Soil Sci. Plant Nutr. 2017;63:553-560.

    Not in our library10.1080/00380768.2017.1395292

  187. 187.

    Konishi N., Saito M., Imagawa F., Kanno K., Yamaya T., Kojima S. Cytosolic glutamine synthetase isozymes play redundant roles in ammonium assimilation under low-ammonium conditions in roots of Arabidopsis thaliana. Plant Cell Physiol. 2018;59:601-613.

    Not in our library10.1093/pcp/pcy014

  188. 188.

    Yamaya T., Kusano M. Evidence supporting distinct functions of three cytosolic glutamine synthetases and two NADH-glutamate synthases in rice. J. Exp. Bot. 2014;65:5519-5525.

    Not in our library10.1093/jxb/eru103

  189. 189.

    Kusano M., Fukushima A., Redestig H., Saito K. Metabolomic approaches toward understanding nitrogen metabolism in plants. J. Exp. Bot. 2011;62:1439-1453.

    Not in our library10.1093/jxb/erq417

  190. 190.

    Forde B.G., Lea P.J. Glutamate in plants: Metabolism, regulation, and signalling. J. Exp. Bot. 2007;58:2339-2358.

    Not in our library10.1093/jxb/erm121

  191. 191.

    Bowsher C.G., Lacey A.E., Hanke G.T., Clarkson D.T., Saker L.R., Stulen I., Emes M.J. The effect of Glc6P uptake and its subsequent oxidation within pea root plastids on nitrite reduction and glutamate synthesis. J. Exp. Bot. 2007;58:1109-1118.

    Not in our library10.1093/jxb/erl269

  192. 192.

    Habash D.Z., Bernard S., Schondelmaier J., Weyen J., Quarrie S.A. The genetics of nitrogen use in hexaploid wheat: N utilisation, development and yield. Theor. Appl. Genet. 2007;114:403-419.

    Not in our library10.1007/s00122-006-0429-5

  193. 193.
    Advances in the functional study of glutamine synthetase in plant abiotic stress tolerance response · 2022
    Not checked
  194. 194.

    Prinsi B., Espen L. Mineral nitrogen sources differently affect root glutamine synthetase isoforms and amino acid balance among organs in maize. BMC Plant Biol. 2015;15:96.

    Not in our library10.1186/s12870-015-0482-9

  195. 195.

    Zhang Z., Xiong S., Wei Y., Meng X., Wang X., Ma X. The role of glutamine synthetase isozymes in enhancing nitrogen use efficiency of N-efficient winter wheat. Sci. Rep. 2017;7:1000.

    Not in our library10.1038/s41598-017-01071-1

  196. 196.

    Habash D., Massiah A., Rong H., Wallsgrove R., Leigh R. The role of cytosolic glutamine synthetase in wheat. Ann. Appl. Biol. 2001;138:83-89.

    Not in our library10.1111/j.1744-7348.2001.tb00087.x

  197. 197.

    Gadaleta A., Nigro D., Marcotuli I., Giancaspro A., Giove S.L., Blanco A. Isolation and characterisation of cytosolic glutamine synthetase (GSe) genes and association with grain protein content in durum wheat. Crop Pasture Sci. 2014;65:38-45.

    Not in our library10.1071/cp13140

  198. 198.

    Tabuchi M., Abiko T., Yamaya T. Assimilation of ammonium ions and reutilization of nitrogen in rice (Oryza sativa L.). J. Exp. Bot. 2007;58:2319-2327.

    Not in our library10.1093/jxb/erm016

  199. 199.

    Ishiyama K., Inoue E., Watanabe-Takahashi A., Obara M., Yamaya T., Takahashi H. Kinetic Properties and Ammonium-dependent Regulation of Cytosolic Isoenzymes of Glutamine Synthetase in Arabidopsis. J. Biol. Chem. 2004;279:16598-16605.

    Not in our library10.1074/jbc.m313710200

  200. 200.
    Influence of Nitrogen and Phosphorus on Microalgal Growth, Biomass, Lipid, and Fatty Acid Production: An Overview · 2021
    No notice
  201. 201.

    Morales-Sánchez D., Martinez-Rodriguez O.A., Kyndt J., Martinez A. Heterotrophic growth of microalgae: Metabolic aspects. World J. Microbiol. Biotechnol. 2015;31:1-9.

    Not in our library10.1007/s11274-014-1773-2

  202. 202.

    Kumar A., Bera S. Revisiting nitrogen utilization in algae: A review on the process of regulation and assimilation. Bioresour. Technol. Rep. 2020;12:100584.

    Not in our library10.1016/j.biteb.2020.100584

  203. 203.

    Vega J.M. Nitrogen and sulfur metabolism in microalgae and plants: 50 years of research. Prog. Bot. 2020;81:1-40.

    Not in our library

  204. 204.

    Zhang H., Zhu J., Gong Z., Zhu J.-K. Abiotic stress responses in plants. Nat. Rev. Genet. 2022;23:104-119.

    Not in our library10.1038/s41576-021-00413-0

  205. 205.

    Sedri M.H., Amini A., Golchin A. Evaluation of Nitrogen Effects on Yield and Drought Tolerance of Rainfed Wheat using Drought Stress Indices. J. Crop Sci. Biotechnol. 2019;22:235-242.

    Not in our library10.1007/s12892-018-0037-0

  206. 206.

    Song J., Wang Y., Pan Y., Pang J., Zhang X., Fan J., Zhang Y. The influence of nitrogen availability on anatomical and physiological responses of Populus alba × P. glandulosa to drought stress. BMC Plant Biol. 2019;19:63.

    Not in our library10.1186/s12870-019-1667-4

  207. 207.

    Xu N., Guo W., Liu J., Du N., Wang R. Increased nitrogen deposition alleviated the adverse effects of drought stress on Quercus variabilis and Quercus mongolica seedlings. Acta Physiol. Plant. 2015;37:107.

    Not in our library10.1007/s11738-015-1853-4

  208. 208.

    Gessler A., Schaub M., McDowell N.G. The role of nutrients in drought-induced tree mortality and recovery. New Phytol. 2017;214:513-520.

    Not in our library10.1111/nph.14340

  209. 209.

    Li S., Zhou L., Addo-Danso S.D., Ding G., Sun M., Wu S., Lin S. Nitrogen supply enhances the physiological resistance of Chinese fir plantlets under polyethylene glycol (PEG)-induced drought stress. Sci. Rep. 2020;10:7509.

    Not in our library10.1038/s41598-020-64161-7

  210. 210.

    Sahay S., Robledo-Arratia L., Glowacka K., Gupta M. Root NRT, NiR, AMT, GS, GOGAT and GDH expression levels reveal NO and ABA mediated drought tolerance in Brassica juncea L. Sci. Rep. 2021;11:7992.

    Not in our library10.1038/s41598-021-86401-0

  211. 211.

    Yang Y., Tang M., Sulpice R., Chen H., Tian S., Ban Y. Arbuscular mycorrhizal fungi alter fractal dimension characteristics of Robinia pseudoacacia L. seedlings through regulating plant growth, leaf water status, photosynthesis, and nutrient concentration under drought stress. J. Plant Growth Regul. 2014;33:612-625.

    Not in our library10.1007/s00344-013-9410-0

  212. 212.

    Khan A., Tan D.K.Y., Afridi M.Z., Luo H., Tung S.A., Ajab M., Fahad S. Nitrogen fertility and abiotic stresses management in cotton crop: A review. Environ. Sci. Pollut. Res. 2017;24:14551-14566.

    Not in our library10.1007/s11356-017-8920-x

  213. 213.

    Diao M., Ma L., Wang J., Cui J., Fu A., Liu H.-y. Selenium Promotes the Growth and Photosynthesis of Tomato Seedlings Under Salt Stress by Enhancing Chloroplast Antioxidant Defense System. J. Plant Growth Regul. 2014;33:671-682.

    Not in our library10.1007/s00344-014-9416-2

  214. 214.

    Zhang N., Zhao B., Zhang H.-J., Weeda S., Yang C., Yang Z.-C., Ren S., Guo Y.-D. Melatonin promotes water-stress tolerance, lateral root formation, and seed germination in cucumber (Cucumis sativus L.). J. Pineal Res. 2013;54:15-23.

    Not in our library10.1111/j.1600-079x.2012.01015.x

  215. 215.

    Yousfi S., Márquez A.J., Betti M., Araus J.L., Serret M.D. Gene expression and physiological responses to salinity and water stress of contrasting durum wheat genotypes. J. Integr. Plant Biol. 2016;58:48-66.

    Not in our library10.1111/jipb.12359

  216. 216.

    Teixeira J., Pereira S. High salinity and drought act on an organ-dependent manner on potato glutamine synthetase expression and accumulation. Environ. Exp. Bot. 2007;60:121-126.

    Not in our library10.1016/j.envexpbot.2006.09.003

  217. 217.

    Cheng L., Wang Y., He Q., Li H., Zhang X., Zhang F. Comparative proteomics illustrates the complexity of drought resistance mechanisms in two wheat (Triticum aestivum L.) cultivars under dehydration and rehydration. BMC Plant Biol. 2016;16:188.

    Not in our library10.1186/s12870-016-0871-8

  218. 218.

    Nagy Z., Németh E., Guóth A., Bona L., Wodala B., Pécsváradi A. Metabolic indicators of drought stress tolerance in wheat: Glutamine synthetase isoenzymes and Rubisco. Plant Physiol. Biochem. 2013;67:48-54.

    Not in our library10.1016/j.plaphy.2013.03.001

  219. 219.

    Chien H.-F., Lin C.C., Wang J.-W., Chen C.T., Kao C.H. Changes in ammonium ion content and glutamine synthetase activity in rice leaves caused by excess cadmium are a consequence of oxidative damage. Plant Growth Regul. 2002;36:41-47.

    Not in our library10.1023/a:1014742014171

  220. 220.

    Balestrasse K.B., Benavides M.P., Gallego S.M., Tomaro M.L. Effect of cadmium stress on nitrogen metabolism in nodules and roots of soybean plants. Funct. Plant Biol. 2003;30:57-64.

    Not in our library10.1071/fp02074

  221. 221.

    Chaffei C., Pageau K., Suzuki A., Gouia H., Ghorbel M.H., Masclaux-Daubresse C. Cadmium Toxicity Induced Changes in Nitrogen Management in Lycopersicon esculentum Leading to a Metabolic Safeguard Through an Amino Acid Storage Strategy. Plant Cell Physiol. 2004;45:1681-1693.

    Not in our library10.1093/pcp/pch192

  222. 222.

    Rana N.K., Mohanpuria P., Yadav S.K. Expression of tea cytosolic glutamine synthetase is tissue specific and induced by cadmium and salt stress. Biol. Plant. 2008;52:361-364.

    Not in our library10.1007/s10535-008-0075-7

  223. 223.

    Kolbert Z., Feigl G., Freschi L., Poór P. Gasotransmitters in action: Nitric oxide-ethylene crosstalk during plant growth and abiotic stress responses. Antioxidants. 2019;8:167.

    Not in our library10.3390/antiox8060167

  224. 224.

    Cao Y., Tanaka K., Nguyen C.T., Stacey G. Extracellular ATP is a central signaling molecule in plant stress responses. Curr. Opin. Plant Biol. 2014;20:82-87.

    Not in our library10.1016/j.pbi.2014.04.009

  225. 225.

    Neill S.J., Desikan R., Hancock J.T. Nitric oxide signalling in plants. New Phytol. 2003;159:11-35.

    Not in our library10.1046/j.1469-8137.2003.00804.x

  226. 226.

    Chamizo-Ampudia A., Sanz-Luque E., Llamas A., Galvan A., Fernandez E. Nitrate reductase regulates plant nitric oxide homeostasis. Trends Plant Sci. 2017;22:163-174.

    Not in our library10.1016/j.tplants.2016.12.001

  227. 227.

    Wilson I.D., Neill S.J., Hancock J.T. Nitric oxide synthesis and signalling in plants. Plant Cell Environ. 2008;31:622-631.

    Not in our library10.1111/j.1365-3040.2007.01761.x

  228. 228.

    Nabi R.B.S., Tayade R., Hussain A., Kulkarni K.P., Imran Q.M., Mun B.-G., Yun B.-W. Nitric oxide regulates plant responses to drought, salinity, and heavy metal stress. Environ. Exp. Bot. 2019;161:120-133.

    Not in our library10.1016/j.envexpbot.2019.02.003

  229. 229.

    Pissolato M.D., Silveira N.M., Prataviera P.J.C., Machado E.C., Seabra A.B., Pelegrino M.T., Sodek L., Ribeiro R.V. Enhanced Nitric Oxide Synthesis Through Nitrate Supply Improves Drought Tolerance of Sugarcane Plants. Front. Plant Sci. 2020;11:970.

    Not in our library10.3389/fpls.2020.00970

  230. 230.

    Procházková D., Sumaira J., Wilhelmová N.a., Pavlíková D., Száková J., Ahmad P., Rasool S. Chapter 11—Reactive Nitrogen Species and the Role of NO in Abiotic Stress. Emerging Technologies and Management of Crop Stress Tolerance. 2014.

    Not in our library

  231. 231.
    Plant Survival in a Changing Environment: The Role of Nitric Oxide in Plant Responses to Abiotic Stress · 2015
    No notice
  232. 232.

    Lau S.-E., Hamdan M.F., Pua T.-L., Saidi N.B., Tan B.C. Plant Nitric Oxide Signaling under Drought Stress. Plants. 2021;10:360.

    Not in our library10.3390/plants10020360

  233. 233.

    Zhang A., Jiang M., Zhang J., Ding H., Xu S., Hu X., Tan M. Nitric oxide induced by hydrogen peroxide mediates abscisic acid-induced activation of the mitogen-activated protein kinase cascade involved in antioxidant defense in maize leaves. New Phytol. 2007;175:36-50.

    Not in our library10.1111/j.1469-8137.2007.02071.x

  234. 234.

    Berger A., Boscari A., Puppo A., Brouquisse R. Nitrate reductases and hemoglobins control nitrogen-fixing symbiosis by regulating nitric oxide accumulation. J. Exp. Bot. 2021;72:873-884.

    Not in our library10.1093/jxb/eraa403

  235. 235.

    Berger A., Boscari A., Horta Araujo N., Maucourt M., Hanchi M., Bernillon S., Rolin D., Puppo A., Brouquisse R. Plant nitrate reductases regulate nitric oxide production and nitrogen-fixing metabolism during the Medicago truncatula–Sinorhizobium meliloti symbiosis. Front. Plant Sci. 2020;11:535004.

    Not in our library10.3389/fpls.2020.01313

  236. 236.

    Wildt J., Kley D., Rockel A., Rockel P., Segschneider H. Emission of NO from several higher plant species. J. Geophys. Res. Atmos. 1997;102:5919-5927.

    Not in our library10.1029/96jd02968

  237. 237.

    Wu Y., Li J., Wang J., Dawuda M.M., Liao W., Meng X., Yuan H., Xie J., Tang Z., Lyu J. Heme is involved in the exogenous ALA-promoted growth and antioxidant defense system of cucumber seedlings under salt stress. BMC Plant Biol. 2022;22:329.

    Not in our library10.1186/s12870-022-03717-3

  238. 238.

    Gan L., Wu X., Zhong Y. Exogenously Applied Nitric Oxide Enhances the Drought Tolerance in Hulless Barley. Plant Prod. Sci. 2015;18:52-56.

    Not in our library10.1626/pps.18.52

  239. 239.

    Ahanger M.A., Qin C., Begum N., Maodong Q., Dong X.X., El-Esawi M., El-Sheikh M.A., Alatar A.A., Zhang L. Nitrogen availability prevents oxidative effects of salinity on wheat growth and photosynthesis by up-regulating the antioxidants and osmolytes metabolism, and secondary metabolite accumulation. BMC Plant Biol. 2019;19:479.

    Not in our library10.1186/s12870-019-2085-3

  240. 240.

    Wang L., Zheng J., Zhou G., Li J., Qian C., Lin G., Li Y., Zuo Q. Moderate nitrogen application improved salt tolerance by enhancing photosynthesis, antioxidants, and osmotic adjustment in rapeseed (Brassica napus L.). Front. Plant Sci. 2023;14:1196319.

    Not in our library10.3389/fpls.2023.1196319

  241. 241.

    Nazir F., Mahajan M., Khatoon S., Albaqami M., Ashfaque F., Chhillar H., Chopra P., Khan M.I.R. Sustaining nitrogen dynamics: A critical aspect for improving salt tolerance in plants. Front. Plant Sci. 2023;14:1087946.

    Not in our library10.3389/fpls.2023.1087946

  242. 242.

    Han M.-L., Lv Q.-Y., Zhang J., Wang T., Zhang C.-X., Tan R.-J., Wang Y.-L., Zhong L.-Y., Gao Y.-Q., Chao Z.-F., et al. Decreasing nitrogen assimilation under drought stress by suppressing DST-mediated activation of Nitrate Reductase 1.2 in rice. Mol. Plant. 2022;15:167-178.

    Not in our library10.1016/j.molp.2021.09.005

  243. 243.

    Hessini K., Issaoui K., Ferchichi S., Saif T., Abdelly C., Siddique K.H.M., Cruz C. Interactive effects of salinity and nitrogen forms on plant growth, photosynthesis and osmotic adjustment in maize. Plant Physiol. Biochem. 2019;139:171-178.

    Not in our library10.1016/j.plaphy.2019.03.005

  244. 244.

    Liu M., Zhao Y., Liu X., Korpelainen H., Li C. Ammonium and nitrate affect sexually different responses to salt stress in Populus cathayana. Physiol. Plant. 2022;174:e13626.

    Not in our library10.1111/ppl.13626

  245. 245.

    Qu C., Liu C., Ze Y., Gong X., Hong M., Wang L., Hong F. Inhibition of Nitrogen and Photosynthetic Carbon Assimilation of Maize Seedlings by Exposure to a Combination of Salt Stress and Potassium-Deficient Stress. Biol. Trace Elem. Res. 2011;144:1159-1174.

    Not in our library10.1007/s12011-011-9037-6

  246. 246.

    Akram M. Effect of Nitrogen Nutrition on Solute Accumulation and Ion Contents of Maize under Sodium Chloride Stress. Commun. Soil Sci. Plant Anal. 2014;45:86-100.

    Not in our library10.1080/00103624.2013.848882

  247. 247.

    Singh M., Singh V.P., Prasad S.M. Responses of photosynthesis, nitrogen and proline metabolism to salinity stress in Solanum lycopersicum under different levels of nitrogen supplementation. Plant Physiol. Biochem. 2016;109:72-83.

    Not in our library10.1016/j.plaphy.2016.08.021

  248. 248.

    Ji Y., Li Q., Liu G., Selvaraj G., Zheng Z., Zou J., Wei Y. Roles of Cytosolic Glutamine Synthetases in Arabidopsis Development and Stress Responses. Plant Cell Physiol. 2019;60:657-671.

    Not in our library10.1093/pcp/pcy235

  249. 249.

    Yan S., Tang Z., Su W., Sun W. Proteomic analysis of salt stress-responsive proteins in rice root. Proteomics. 2005;5:235-244.

    Not in our library10.1002/pmic.200400853

  250. 250.

    Zhao F.G., Qin P. Protective effect of exogenous polyamines on root tonoplast function against salt stress in barley seedlings. Plant Growth Regul. 2004;42:97-103.

    Not in our library10.1023/b:grow.0000017478.40445.bc

  251. 251.

    Hasanuzzaman M., Oku H., Nahar K., Bhuyan M.H.M.B., Mahmud J.A., Baluska F., Fujita M. Nitric oxide-induced salt stress tolerance in plants: ROS metabolism, signaling, and molecular interactions. Plant Biotechnol. Rep. 2018;12:77-92.

    Not in our library10.1007/s11816-018-0480-0

  252. 252.

    Bhardwaj S., Kapoor D., Singh S., Gautam V., Dhanjal D.S., Jan S., Ramamurthy P.C., Prasad R., Singh J. Nitric Oxide: A Ubiquitous Signal Molecule for Enhancing Plant Tolerance to Salinity Stress and Their Molecular Mechanisms. J. Plant Growth Regul. 2021;40:2329-2341.

    Not in our library10.1007/s00344-021-10394-3

  253. 253.
    Nitric Oxide Mitigates Salt Stress by Regulating Levels of Osmolytes and Antioxidant Enzymes in Chickpea · 2016
    No notice
  254. 254.

    Zheng C., Jiang D., Liu F., Dai T., Liu W., Jing Q., Cao W. Exogenous nitric oxide improves seed germination in wheat against mitochondrial oxidative damage induced by high salinity. Environ. Exp. Bot. 2009;67:222-227.

    Not in our library10.1016/j.envexpbot.2009.05.002

  255. 255.

    Lopez G., Ahmadi S.H., Amelung W., Athmann M., Ewert F., Gaiser T., Gocke M.I., Kautz T., Postma J., Rachmilevitch S., et al. Nutrient deficiency effects on root architecture and root-to-shoot ratio in arable crops. Front. Plant Sci. 2023;13:1067498.

    Not in our library10.3389/fpls.2022.1067498

  256. 256.

    Falcioni R., Moriwaki T., Rodrigues M., de Oliveira K.M., Furlanetto R.H., dos Reis A.S., dos Santos G.L.A.A., Mendonça W.A., Crusiol L.G.T., Gonçalves J.V.F., et al. Nutrient deficiency lowers photochemical and carboxylation efficiency in tobacco. Theor. Exp. Plant Physiol. 2023;35:81-97.

    Not in our library10.1007/s40626-023-00268-2

  257. 257.

    Sattelmacher B., Addiscott T.M. AP Whitmore and DS Powlson: Farming, Fertilizers and the Nitrate Problem. 1991.

    Not in our library

  258. 258.

    Aparicio V., Costa J.L., Zamora M. Nitrate leaching assessment in a long-term experiment under supplementary irrigation in humid Argentina. Agric. Water Manag. 2008;95:1361-1372.

    Not in our library10.1016/j.agwat.2008.06.003

  259. 259.

    Ward M.H. Too Much of a Good Thing? Nitrate from Nitrogen Fertilizers and Cancer. Rev. Environ. Health. 2009;24:357-363.

    Not in our library10.1515/reveh.2009.24.4.357

  260. 260.

    Ju X.T., Kou C.L., Zhang F.S., Christie P. Nitrogen balance and groundwater nitrate contamination: Comparison among three intensive cropping systems on the North China Plain. Environ. Pollut. 2006;143:117-125.

    Not in our library10.1016/j.envpol.2005.11.005

  261. 261.

    Maghanga J.K., Kituyi J.L., Kisinyo P.O., Ng’etich W.K. Impact of Nitrogen Fertilizer Applications on Surface Water Nitrate Levels within a Kenyan Tea Plantation. J. Chem. 2013;2013:196516.

    Not in our library10.1155/2013/196516

  262. 262.

    Bryan N.S., van Grinsven H., Sparks D.L. Chapter Three—The Role of Nitrate in Human Health. Advances in Agronomy. 2013;Volume 119:153-182.

    Not in our library

  263. 263.

    Fraser P., Chilvers C., Beral V., Hill M.J. Nitrata and Human Cancer: A Review of the Evidence. Int. J. Epidemiol. 1980;9:3-12.

    Not in our library10.1093/ije/9.1.3

  264. 264.
    Blue babies and nitrate-contaminated well water. · 2000
    No notice
  265. 265.

    Fischer R.A., Connor D.J. Issues for cropping and agricultural science in the next 20 years. Field Crops Res. 2018;222:121-142.

    Not in our library10.1016/j.fcr.2018.03.008

  266. 266.

    Liu Q., Chen X., Wu K., Fu X. Nitrogen signaling and use efficiency in plants: What’s new? Curr. Opin. Plant Biol. 2015;27:192-198.

    Not in our library10.1016/j.pbi.2015.08.002

  267. 267.

    Xu G., Fan X., Miller A.J. Plant Nitrogen Assimilation and Use Efficiency. Annu. Rev. Plant Biol. 2012;63:153-182.

    Not in our library10.1146/annurev-arplant-042811-105532

  268. 268.

    Motavalli P.P., Goyne K.W., Udawatta R.P. Environmental Impacts of Enhanced-Efficiency Nitrogen Fertilizers. Crop Manag. 2008;7:1-15.

    Not in our library10.1094/cm-2008-0730-02-rv

  269. 269.

    Woods J., Williams A., Hughes J.K., Black M., Murphy R. Energy and the food system. Philos. Trans. R. Soc. B Biol. Sci. 2010;365:2991-3006.

    Not in our library10.1098/rstb.2010.0172

  270. 270.

    Ntatsi G., Karkanis A., Yfantopoulos D., Pappa V., Konosonoka I.H., Travlos I., Bilalis D., Bebeli P., Savvas D. Evaluation of the field performance, nitrogen fixation efficiency and competitive ability of pea landraces grown under organic and conventional farming systems. Arch. Agron. Soil Sci. 2019;65:294-307.

    Not in our library10.1080/03650340.2018.1501155

  271. 271.
    Nitrogen fixation in a landrace of maize is supported by a mucilage-associated diazotrophic microbiota · 2018
    No notice
  272. 272.

    Barnawal D., Maji D., Bharti N., Chanotiya C.S., Kalra A. ACC Deaminase-Containing Bacillus subtilis Reduces Stress Ethylene-Induced Damage and Improves Mycorrhizal Colonization and Rhizobial Nodulation in Trigonella foenum-graecum Under Drought Stress. J. Plant Growth Regul. 2013;32:809-822.

    Not in our library10.1007/s00344-013-9347-3

  273. 273.

    Kour D., Rana K.L., Yadav A.N., Yadav N., Kumar M., Kumar V., Vyas P., Dhaliwal H.S., Saxena A.K. Microbial biofertilizers: Bioresources and eco-friendly technologies for agricultural and environmental sustainability. Biocatal. Agric. Biotechnol. 2020;23:101487.

    Not in our library10.1016/j.bcab.2019.101487

  274. 274.

    Compant S., Cambon M.C., Vacher C., Mitter B., Samad A., Sessitsch A. The plant endosphere world—Bacterial life within plants. Environ. Microbiol. 2021;23:1812-1829.

    Not in our library10.1111/1462-2920.15240

  275. 275.

    Rane N.R., Tapase S., Kanojia A., Watharkar A., Salama E.-S., Jang M., Kumar Yadav K., Amin M.A., Cabral-Pinto M.M.S., Jadhav J.P., et al. Molecular insights into plant–microbe interactions for sustainable remediation of contaminated environment. Bioresour. Technol. 2022;344:126246.

    Not in our library10.1016/j.biortech.2021.126246

  276. 276.

    Dubois O. The State of the World’s Land and Water Resources for Food and Agriculture: Managing Systems at Risk. 2011.

    Not in our library

  277. 277.

    Francis B., Aravindakumar C.T., Brewer P.B., Simon S. Plant nutrient stress adaptation: A prospect for fertilizer limited agriculture. Environ. Exp. Bot. 2023;213:105431.

    Not in our library10.1016/j.envexpbot.2023.105431

  278. 278.

    Hartemink A.E. The definition of soil since the early 1800s. Adv. Agron. 2016;137:73-126.

    Not in our library

  279. 279.

    Campanhola C., Pandey S. Sustainable Food and Agriculture: An Integrated Approach. 2018.

    Not in our library

  280. 280.

    Igiehon N.O., Babalola O.O. Biofertilizers and sustainable agriculture: Exploring arbuscular mycorrhizal fungi. Appl. Microbiol. Biotechnol. 2017;101:4871-4881.

    Not in our library10.1007/s00253-017-8344-z

  281. 281.

    Bodelier P.L. Toward understanding, managing, and protecting microbial ecosystems. Front. Microbiol. 2011;2:80.

    Not in our library10.3389/fmicb.2011.00080

  282. 282.

    Bhattarai A., Bhattarai B., Pandey S. Variation of soil microbial population in different soil horizons. J. Microbiol. Exp. 2015;2:00044.

    Not in our library10.15406/jmen.2015.02.00044

  283. 283.
    The structure and function of the global citrus rhizosphere microbiome · 2018
    No notice
  284. 284.
    Probiotic Diversity Enhances Rhizosphere Microbiome Function and Plant Disease Suppression · 2016
    No notice
  285. 285.

    Mendes R., Garbeva P., Raaijmakers J.M. The rhizosphere microbiome: Significance of plant beneficial, plant pathogenic, and human pathogenic microorganisms. FEMS Microbiol. Rev. 2013;37:634-663.

    Not in our library10.1111/1574-6976.12028

  286. 286.

    Mitter E.K., Tosi M., Obregón D., Dunfield K.E., Germida J.J. Rethinking crop nutrition in times of modern microbiology: Innovative biofertilizer technologies. Front. Sustain. Food Syst. 2021;5:606815.

    Not in our library10.3389/fsufs.2021.606815

  287. 287.

    Nannipieri P., Ascher-Jenull J., Ceccherini M.T., Pietramellara G., Renella G., Schloter M. Beyond microbial diversity for predicting soil functions: A mini review. Pedosphere. 2020;30:5-17.

    Not in our library10.1016/s1002-0160(19)60824-6

  288. 288.

    Johansson J.F., Paul L.R., Finlay R.D. Microbial interactions in the mycorrhizosphere and their significance for sustainable agriculture. FEMS Microbiol. Ecol. 2004;48:1-13.

    Not in our library10.1016/j.femsec.2003.11.012

  289. 289.

    Saccá M.L., Barra Caracciolo A., Di Lenola M., Grenni P. Ecosystem services provided by soil microorganisms. Soil Biological Communities and Ecosystem Resilience. 2017;9-24.

    Not in our library

  290. 290.

    Nannipieri P., Ascher J., Ceccherini M., Landi L., Pietramellara G., Renella G. Microbial diversity and soil functions. Eur. J. Soil Sci. 2003;54:655-670.

    Not in our library10.1046/j.1351-0754.2003.0556.x

  291. 291.

    Masson-Boivin C., Giraud E., Perret X., Batut J. Establishing nitrogen-fixing symbiosis with legumes: How many rhizobium recipes? Trends Microbiol. 2009;17:458-466.

    Not in our library10.1016/j.tim.2009.07.004

  292. 292.

    Mendoza-Suárez M., Andersen S.U., Poole P.S., Sánchez-Cañizares C. Competition, nodule occupancy, and persistence of inoculant strains: Key factors in the rhizobium-legume symbioses. Front. Plant Sci. 2021;12:690567.

    Not in our library10.3389/fpls.2021.690567

  293. 293.

    Holland P.L. Introduction: Reactivity of Nitrogen from the Ground to the Atmosphere. Chem. Rev. 2020;120:4919-4920.

    Not in our library10.1021/acs.chemrev.0c00361

  294. 294.

    Nyaga J.W., Njeru E.M. Potential of native rhizobia to improve cowpea growth and production in semiarid regions of Kenya. Front. Agron. 2020;2:606293.

    Not in our library10.3389/fagro.2020.606293

  295. 295.

    Yamada Y., Hoshino K.-i., Ishikawa T. The phylogeny of acetic acid bacteria based on the partial sequences of 16S ribosomal RNA: The elevation of the subgenus Gluconoacetobacter to the generic level. Biosci. Biotechnol. Biochem. 1997;61:1244-1251.

    Not in our library10.1271/bbb.61.1244

  296. 296.

    Yan D., Tajima H., Cline L.C., Fong R.Y., Ottaviani J.I., Shapiro H.Y., Blumwald E. Genetic modification of flavone biosynthesis in rice enhances biofilm formation of soil diazotrophic bacteria and biological nitrogen fixation. Plant Biotechnol. J. 2022;20:2135-2148.

    Not in our library10.1111/pbi.13894

  297. 297.

    Cesco S., Mimmo T., Tonon G., Tomasi N., Pinton R., Terzano R., Neumann G., Weisskopf L., Renella G., Landi L. Plant-borne flavonoids released into the rhizosphere: Impact on soil bio-activities related to plant nutrition. A review. Biol. Fertil. Soils. 2012;48:123-149.

    Not in our library10.1007/s00374-011-0653-2

  298. 298.

    Wang D., Xu A., Elmerich C., Ma L.Z. Biofilm formation enables free-living nitrogen-fixing rhizobacteria to fix nitrogen under aerobic conditions. ISME J. 2017;11:1602-1613.

    Not in our library10.1038/ismej.2017.30

  299. 299.

    Flemming H.-C., Wingender J., Szewzyk U., Steinberg P., Rice S.A., Kjelleberg S. Biofilms: An emergent form of bacterial life. Nat. Rev. Microbiol. 2016;14:563-575.

    Not in our library10.1038/nrmicro.2016.94

  300. 300.

    Young J.M., Leschine S.B., Reguera G. Reversible control of biofilm formation by Cellulomonas spp. in response to nitrogen availability. Environ. Microbiol. 2012;14:594-604.

    Not in our library10.1111/j.1462-2920.2011.02596.x

  301. 301.

    Pankievicz V., Irving T.B., Maia L.G., Ané J.-M. Are we there yet? The long walk towards the development of efficient symbiotic associations between nitrogen-fixing bacteria and non-leguminous crops. BMC Biol. 2019;17:1-17.

    Not in our library10.1186/s12915-019-0710-0

  302. 302.

    Ike A., Sriprang R., Ono H., Murooka Y., Yamashita M. Bioremediation of cadmium contaminated soil using symbiosis between leguminous plant and recombinant rhizobia with the MTL4 and the PCS genes. Chemosphere. 2007;66:1670-1676.

    Not in our library10.1016/j.chemosphere.2006.07.058

  303. 303.
    Nitrogen Fixation in Cereals · 2018
    No notice
  304. 304.

    Hill S., Kennedy C., Kavanagh E., Goldberg R.B., Hanau R. Nitrogen fixation gene (nifL) involved in oxygen regulation of nitrogenase synthesis in K. peumoniae. Nature. 1981;290:424-426.

    Not in our library10.1038/290424a0

  305. 305.

    Dixon R., Kahn D. Genetic regulation of biological nitrogen fixation. Nat. Rev. Microbiol. 2004;2:621-631.

    Not in our library10.1038/nrmicro954

  306. 306.

    Burgess B.K., Lowe D.J. Mechanism of molybdenum nitrogenase. Chem. Rev. 1996;96:2983-3012.

    Not in our library10.1021/cr950055x

  307. 307.

    Bjornsson R., Delgado-Jaime M.U., Lima F.A., Sippel D., Schlesier J., Weyhermüller T., Einsle O., Neese F., DeBeer S. Molybdenum L-Edge XAS Spectra of MoFe Nitrogenase. Z. Für Anorg. Und Allg. Chem. 2015;641:65-71.

    Not in our library10.1002/zaac.201400446

  308. 308.

    Setubal J.C., Dos Santos P., Goldman B.S., Ertesvaåg H., Espin G., Rubio L.M., Valla S., Almeida N.F., Balasubramanian D., Cromes L. Genome sequence of Azotobacter vinelandii, an obligate aerobe specialized to support diverse anaerobic metabolic processes. J. Bacteriol. 2009;191:4534-4545.

    Not in our library10.1128/jb.00504-09

  309. 309.

    Yan Y., Yang J., Dou Y., Chen M., Ping S., Peng J., Lu W., Zhang W., Yao Z., Li H. Nitrogen fixation island and rhizosphere competence traits in the genome of root-associated Pseudomonas stutzeri A1501. Proc. Natl. Acad. Sci. USA. 2008;105:7564-7569.

    Not in our library10.1073/pnas.0801093105

  310. 310.

    Lalucat J., Bennasar A., Bosch R., García-Valdés E., Palleroni N.J. Biology of Pseudomonas stutzeri. Microbiol. Mol. Biol. Rev. 2006;70:510-547.

    Not in our library10.1128/mmbr.00047-05

  311. 311.

    Dingler C., Kuhla J., Wassink H., Oelze J. Levels and activities of nitrogenase proteins in Azotobacter vinelandii grown at different dissolved oxygen concentrations. J. Bacteriol. 1988;170:2148-2152.

    Not in our library10.1128/jb.170.5.2148-2152.1988

  312. 312.

    Desnoues N., Lin M., Guo X., Ma L., Carreño-Lopez R., Elmerich C. Nitrogen fixation genetics and regulation in a Pseudomonas stutzeri strain associated with rice. Microbiology. 2003;149:2251-2262.

    Not in our library10.1099/mic.0.26270-0

  313. 313.

    Oelze J. Respiratory protection of nitrogenase in Azotobacter species: Is a widely held hypothesis unequivocally supported by experimental evidence? FEMS Microbiol. Rev. 2000;24:321-333.

    Not in our library10.1111/j.1574-6976.2000.tb00545.x

  314. 314.

    Curatti L., Brown C.S., Ludden P.W., Rubio L.M. Genes required for rapid expression of nitrogenase activity in Azotobacter vinelandii. Proc. Natl. Acad. Sci. USA. 2005;102:6291-6296.

    Not in our library10.1073/pnas.0501216102

  315. 315.

    Kelly M., Poole R., Yates M., Kennedy C. Cloning and mutagenesis of genes encoding the cytochrome bd terminal oxidase complex in Azotobacter vinelandii: Mutants deficient in the cytochrome d complex are unable to fix nitrogen in air. J. Bacteriol. 1990;172:6010-6019.

    Not in our library10.1128/jb.172.10.6010-6019.1990

  316. 316.

    Dedysh S.N., Ricke P., Liesack W. NifH and NifD phylogenies: An evolutionary basis for understanding nitrogen fixation capabilities of methanotrophic bacteria. Microbiology. 2004;150:1301-1313.

    Not in our library10.1099/mic.0.26585-0

  317. 317.

    Raymond J., Siefert J.L., Staples C.R., Blankenship R.E. The natural history of nitrogen fixation. Mol. Biol. Evol. 2004;21:541-554.

    Not in our library10.1093/molbev/msh047

  318. 318.

    Henson B.J., Watson L.E., Barnum S.R. The evolutionary history of nitrogen fixation, as assessed by nif D. J. Mol. Evol. 2004;58:390-399.

    Not in our library10.1007/s00239-003-2560-0

  319. 319.

    Romanovsky V.E., Drozdov D., Oberman N.G., Malkova G., Kholodov A.L., Marchenko S., Moskalenko N.G., Sergeev D., Ukraintseva N., Abramov A. Thermal state of permafrost in Russia. Permafr. Periglac. Process. 2010;21:136-155.

    Not in our library10.1002/ppp.683

  320. 320.

    Christiansen H.H., Etzelmüller B., Isaksen K., Juliussen H., Farbrot H., Humlum O., Johansson M., Ingeman-Nielsen T., Kristensen L., Hjort J. The thermal state of permafrost in the nordic area during the international polar year 2007–2009. Permafr. Periglac. Process. 2010;21:156-181.

    Not in our library10.1002/ppp.687

  321. 321.

    Grosse G., Goetz S., McGuire A.D., Romanovsky V.E., Schuur E.A. Changing permafrost in a warming world and feedbacks to the Earth system. Environ. Res. Lett. 2016;11:040201.

    Not in our library10.1088/1748-9326/11/4/040201

  322. 322.

    Hugelius G., Strauss J., Zubrzycki S., Harden J.W., Schuur E.A., Ping C.-L., Schirrmeister L., Grosse G., Michaelson G.J., Koven C.D. Estimated stocks of circumpolar permafrost carbon with quantified uncertainty ranges and identified data gaps. Biogeosciences. 2014;11:6573-6593.

    Not in our library10.5194/bg-11-6573-2014

  323. 323.

    Schuur E.A., McGuire A.D., Schädel C., Grosse G., Harden J.W., Hayes D.J., Hugelius G., Koven C.D., Kuhry P., Lawrence D.M. Climate change and the permafrost carbon feedback. Nature. 2015;520:171-179.

    Not in our library10.1038/nature14338

  324. 324.

    Schädel C., Bader M.K.-F., Schuur E.A., Biasi C., Bracho R., Čapek P., De Baets S., Diáková K., Ernakovich J., Estop-Aragones C. Potential carbon emissions dominated by carbon dioxide from thawed permafrost soils. Nat. Clim. Chang. 2016;6:950-953.

    Not in our library10.1038/nclimate3054

  325. 325.

    Harden J.W., Koven C.D., Ping C.L., Hugelius G., McGuire A.D., Camill P., Jorgenson T., Kuhry P., Michaelson G.J., O’Donnell J.A. Field information links permafrost carbon to physical vulnerabilities of thawing. Geophys. Res. Lett. 2012;39:L15704.

    Not in our library10.1029/2012gl051958

  326. 326.
    Nitrous oxide emissions from soils: how well do we understand the processes and their controls? · 2013
    No notice
  327. 327.

    Kuypers M.M., Marchant H.K., Kartal B. The microbial nitrogen-cycling network. Nat. Rev. Microbiol. 2018;16:263-276.

    Not in our library10.1038/nrmicro.2018.9

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