Control of the rhizobium–legume symbiosis by the plant nitrogen demand is tightly integrated at the whole plant level and requires inter-organ systemic signaling

Abstract

Symbiotic nodules formed on legume roots with rhizobia fix atmospheric N₂. Bacteria reduce N₂ to NH₄⁺ that is assimilated into amino acids by the plant. In return, the plant provides photosynthates to fuel the symbiotic nitrogen fixation. Symbiosis is tightly adjusted to the whole plant nutritional demand and to the plant photosynthetic capacities, but regulatory circuits behind this control remain poorly understood. The use of split-root systems combined with biochemical, physiological, metabolomic, transcriptomic, and genetic approaches revealed that multiple pathways are acting in parallel. Systemic signaling mechanisms of the plant N demand are required for the control of nodule organogenesis, mature nodule functioning, and nodule senescence. N-satiety/N-deficit systemic signaling correlates with rapid variations of the nodules’ sugar levels, tuning symbiosis by C resources allocation. These mechanisms are responsible for the adjustment of plant symbiotic capacities to the mineral N resources. On the one hand, if mineral N can satisfy the plant N demand, nodule formation is inhibited, and nodule senescence is activated. On the other hand, local conditions (abiotic stresses) may impair symbiotic activity resulting in plant N limitation. In these conditions, systemic signaling may compensate the N deficit by stimulating symbiotic root N foraging. In the past decade, several molecular components of the systemic signaling pathways controlling nodule formation have been identified, but a major challenge remains, that is, to understand their specificity as compared to the mechanisms of non-symbiotic plants that control root development and how they contribute to the whole plant phenotypes. Less is known about the control of mature nodule development and functioning by N and C nutritional status of the plant, but a hypothetical model involving the sucrose allocation to the nodule as a systemic signaling process, the oxidative pentose phosphate pathway, and the redox status as potential effectors of this signaling is emerging. This work highlights the importance of organism integration in plant biology.

Introduction

Nitrate (NO₃⁻) and ammonium (NH₄⁺) are the major forms of inorganic nitrogen (N) in the soil. However, plant growth in terrestrial ecosystems is often limited by N availability (Verhoeven et al., 1996; Ågren et al., 2012). Approximately 65 million years ago, plants of the legume family (Fabaceae) and soil bacteria of the Rhizobia type gain the capacity to establish a symbiosis whose function is to reduce atmospheric nitrogen (N₂) to ammonia (NH₃/NH₄⁺) within the bacteria, and then transfer the NH₄⁺ to the plant when its N demand is not satisfied by mineral N present in the soil (Roy et al., 2020). Although atmospheric N₂ is a non-limiting N resource, symbiotic nitrogen fixation (SNF) generally does not entirely meet the plant’s N requirements. Indeed, N acquisition through SNF generally does not reach the same level as the uptake of NO₃⁻ and NH₄⁺ when these ions are present in non-limiting concentrations (Ruffel et al., 2008). However, symbiosis allows legume holobionts (i.e., plant in association with its symbiotic bacteria) to grow on poor soils lacking inorganic N. Nevertheless, when enough mineral N is present in soils, legume symbiosis is inhibited and plants satisfy their N demand by mineral N acquisition as non-symbiotic higher plants.

SNF takes place in a new organ, the nodule (Figure 1), in which the plant hosts and nourishes the bacteria (Oldroyd et al., 2011). After a stage of mutual recognition of the two partners, involving plant flavonoids and bacterial lipochito-oligosaccharides (the Nod factors), the bacteria penetrate inside the root hairs via a specific structure, the infection thread, while the root cortical cells divide to initiate nodule formation (Roy et al., 2020). The infection thread grows and crosses the root hair and then the cortical cells to reach the cells of the young growing nodule inside which the bacteria are released. In the nodule, the bacteria differentiate into bacteroids and acquire the ability to reduce N₂ to NH₃/NH₄⁺ through the activity of a specific enzyme, the nitrogenase (Oldroyd and Downie, 2008). The nodules are of either indeterminate (clover, Medicago, alfalfa, and pea) or determinate (soybean, cowpea, and bean) type (Hirsch, 1992). Indeterminate nodules have a persistent meristem and are composed of four distinct zones (Figure 1): zone I (meristematic zone) where cells divide, zone II (infection zone) where bacteria infect cells of the plant and differentiate into bacteroids (i.e., specialized terminally differentiated bacteria unable to divide anymore), zone III (fixation zone) where the bacteroids reduce N₂ to NH₃/NH₄⁺, and zone IV (senescence zone) where plant cells and bacteroids enter in senescence (Timmers et al., 2000). The determinate nodules have no persistent meristem and develop by cell expansion. Reduction of N₂ by nitrogenase and subsequent transfer of NH₄⁺ to the plant partner is the central process of symbiosis (Figure 2; Oldroyd and Downie, 2008). As nitrogenase is irreversibly inhibited by traces of oxygen (O₂), the concentration of O₂ inside the nodules is very low, approximately 10 to 40 nM (Appleby, 1992). Thus, the nodules must move from a normoxic environment, at the beginning of their development, to a microoxic one in the fixation zone of mature N₂-fixing nodules. In exchange for reduced N, the plant provides carbon (C) in the form of dicarboxylic acids to the bacterial partner (Yurgel and Kahn, 2004; Udvardi and Poole, 2013). Terminally differentiated bacteroids display a metabolic specialization in nitrogen fixation. On the one hand, repression of NH₄⁺ assimilation through the Glutamine synthetase/Glutamate synthase (GS/GOGAT) cycle makes the bacteroids dependent on amino acids supplied by the plant (Patriarca et al., 2002; Prell et al., 2009; Oldroyd et al., 2011). On the other hand, NH₄⁺ produced by nitrogenase is exported outside of the bacteroid, acquired, and actively assimilated in surrounding plant cells in the presence of carbon skeleton acceptors derived from photosynthates translocated from shoots to roots and nodules (Figure 2).

Figure 1

Figure 2

The N supply to the plant by nodules requires various regulatory mechanisms operating through the interplay between C and N metabolisms, the sensing of the whole plant N status, the adjustment of nodule capacity to the plant N demand, the supply of O₂ to the nodules, and/or the redox homeostasis (Figure 2; Schulze, 2004; Schwember et al., 2019; Lindström and Mousavi, 2020; Chaulagain and Frugoli, 2021). Over the last 15 years, multiple regulatory mechanisms, acting either locally in response to the nodule environment or at the whole symbiotic plant level and involving systemic signaling, have been evidenced. This review aims to summarize our current knowledge on the regulation of the N-fixing symbiosis by the plant N and C nutritional status through the interplay of N, C, and energy metabolisms, and the diverse local and systemic signaling mechanisms. The current knowledge of the various pathways characterized at the molecular and genetic levels will be discussed regarding their impact on the whole plant phenotype. Finally, we will focus on the future challenges toward the understanding of the control of symbiosis by the plant nutritional demand and the attempt to understand crosstalk, interplays, and emerging properties of symbiotic holobionts in the context of a fluctuating environment. Oxygen plays a key role in regulating nitrogen fixation and nodule energy metabolism (Schulze, 2004; Schwember et al., 2019; Booth et al., 2021; Schulte et al., 2021). This topic is not exhaustively reviewed in this paper but will be occasionally mentioned when a crosstalk with regulation of symbiosis by the plant nutritional demand is suggested.

Symbiotic capacities are often limited by the environment or restricted by the plant

Rhizobium–legume symbiosis may be seen as an adaptive response of the legume–rhizobium holobiont to circumvent plant N deficit by activating a new N acquisition pathway from air, an unlimited N source (Figure 2). However, legume plants relying on N₂ fixation are frequently N limited, indicating that symbiosis may not be sufficient to fulfill alone the N requirements for plant optimal nitrogen nutrition (Crozat et al., 1994; Gan et al., 2002; Moreau et al., 2008). Multiple factors contribute to the limitation of symbiotic capacity.

A major cause of this limitation is the carbon cost of the nitrogen fixation process (Minchin and Witty, 2005). It is generally observed that symbiotic development is tightly correlated to the plant capacity to supply the symbiotic structures with carbon (C) metabolites required for its formation, persistence, and functioning (Walsh et al., 1987; Voisin et al., 2003). Although nodules represent a small part of the plant mass, they can consume more than 25% of the products of photosynthesis for SNF (Schuize et al., 1999; Vance, 2008). The carbon cost per unit of fixed N (g C per g N fixed) was shown to vary widely with species, growth stage, and environmental conditions, ranging from 1.4 to 12 g C per gram fixed N (Schwember et al., 2019). The limitation of symbiosis by photosynthesis and the supply of carbon to the nodules has been a matter of debate. On one hand, some authors have argued that under normal growth conditions (non-limiting water and mineral supply, optimal photoperiod, and light intensity), the supply of sugars from photosynthesis to the nodules may not be limiting (Vance and Heichel, 1991; Schulze, 2004). On the other hand, under environmental stress, such as water deficiency, a reduced availability of C for bacteroid respiration and nitrogenase activity was associated with the decline in N₂ fixation (Gordon et al., 1999; Baier et al., 2007). Furthermore, several studies have shown that elevated CO₂ concentrations (eCO₂) stimulate N₂ fixation and plant biomass production, demonstrating that photosynthesis is effectively limiting symbiosis (Rogers et al., 2006; Sanz-Sáez et al., 2010; Lam et al., 2012; Li et al., 2017; Parvin et al., 2020). In M. truncatula under eCO₂, nodule number and size are increased and most N₂ fixation-related genes are upregulated (Guo et al., 2013). This was further confirmed by Parvin et al. (2020) who showed, either in normal growth condition or under hydric stress, that faba bean under eCO₂ is only able to increase its C gain if nodule activity is maintained. This response of symbiotic legumes to eCO₂ is original as compared to non-symbiotic C3 plants supplied by NO₃⁻ as N source displaying an eCO₂ acclimation and a reduction of NO₃⁻ uptake and assimilation (Stitt and Krapp, 1999; Rogers et al., 2006; Guo et al., 2013). The causes of this acclimation to eCO₂ are not yet understood, but several hypotheses have been recently raised, such as a lower NO₃⁻ concentration in most plant organs, a reduced NO₃⁻ acquisition due to a decreased leaf transpiration, an insufficient NADH power for NO₃⁻ reduction due to reduced photorespiration under eCO₂, or the repression of most NO₃⁻ uptake and assimilation systems by eCO₂ (Gojon et al., 2022).

Nevertheless, C supply is not the only cause of the limitation of symbiotic capacities. Firstly, the nodule development process in N-limited plants requires several days to result to active SNF. Severe N deficit during this lag period is often detrimental for plant growth (especially if the N demand of the young plant is high) and may inhibit the process (Moreau et al., 2008). Secondly, as soon as the symbiosis is established, the SNF efficiency is frequently not at its maximum. Compatible rhizobia forming natural populations in the soil, able to form nodule with a legume host, may result in contrasted levels of SNF (Laguerre et al., 2012; Bourion et al., 2017; Boivin et al., 2020). Thirdly, symbiotic organs are highly sensitive to local environmental abiotic constraints such as drought, heavy metal, temperature, soil pH, or mineral deficiencies (phosphorus, sulfur) that may drastically inhibit SNF (Durand et al., 1987; Liu et al., 2011; Ferguson et al., 2013; Marino et al., 2013; Gil-Quintana et al., 2013b). The dynamics and the fluctuation of these constraints, as well as the time required to establish the new symbiotic structures necessary for the plant N limitation recovery, must also be considered. Fourthly, the nodule proliferation is tightly controlled and generally limited by the plant at multiple steps of the nodule development. Split-root studies have been used to characterize the systemic control of symbiosis by the whole plant (Figure 3). The autoregulation of nodule number (AON) mechanism limits the new infections by compatible rhizobia as soon as a first infection wave is progressing toward the formation of the symbiotic organ (Kosslak et al., 1983; Kosslak and Bohlool, 1984; Mathews et al., 1989; Olsson et al., 1989; Reid et al., 2011b; Kassaw et al., 2015; Ferguson et al., 2019). The inhibition occurs rapidly after infection before the completion of organogenesis and the activation of nitrogen fixation activity. The formation of spontaneous pseudo-nodules or nodules induced by R. meliloti mutants defective in their ability to invade and multiply within host tissues elicits the AON-related feedback suppression of nodule formation similarly to that elicited by the wild-type bacteria (Caetano-Anollés et al., 1990; Caetano-Anollés et al., 1991). Therefore, AON is related to developmental rather than to nutritional feedback. However, AON is also regulated by the plant N demand. In response to N deficit, the symbiotic plant releases the AON repression to increase its nodule number (Figure 3A; Jeudy et al., 2010; Laguerre et al., 2012). Fifthly, when the symbiotic organ is formed, its behavior remained tightly controlled by the plant. At the nodule level, the absence of N₂ fixation triggers a local “plant sanction” response associated with the rapid arrest of nodule growth and the reduction of cultivable bacteria in the nodule (Kiers et al., 2003; Oono et al., 2009; Oono et al., 2011). Mature nodule development is tightly controlled by the systemic signaling of the plant nitrogen demand especially in indeterminate legumes (Figure 3B). On one hand, the plant N deficit stimulates the nodule expansion (as well as bacteroid differentiation), resulting in an increase in N₂ fixation activity (Jeudy et al., 2010; Laguerre et al., 2012). On the other hand, the plant N satiety activates the destruction of bacteroids and the senescence of the organ (Pérez Guerra et al., 2010; Lambert et al., 2020).

Figure 3

Ruffel et al. (2008); Jeudy et al. (2010); Laguerre et al. (2012); Lambert et al. (2020), and Pervent et al. (2021). M. truncatula plants are cultivated hydroponically. The root system of each plant is separated into two compartments. Each compartment receives specific N supplies: either low level of mineral N supply (0.5 mM KNO₃), high level of mineral N supply, air (N₂/O₂ 80/20 v/v), or no gaseous N supply (Ar/O₂ 80/20 v/v). At the beginning of the experiment, a contrasted N supply is applied to a half-root system. The N treatment modifies the whole plant N status and the whole plant N demand without affecting the environment and the untreated half-root system. Response of the untreated roots to this treatment results necessarily from systemic signaling originated from the rest of plant to these roots. (A) Split root systems used to study the control of nodule formation by plant N demand (Pervent et al., 2021). Non symbiotic plants are used. Contrasted mineral N supply applied to half-root systems may either fully satisfy the plant N demand (N satiety) or results in plant N limitation (N deficit). The response to inoculation by rhizobium in untreated roots strongly relies on the level of N supply of the plant revealing the strong systemic control of nodule formation by systemic signaling of the plant N demand. (B) Split root systems used to study the control of mature nodules by the plant N demand (Jeudy et al., 2010; Laguerre et al., 2012; Lambert et al., 2020). Symbiotic plants supplied with air as a unique N source through the function of mature N₂ fixing nodules are used. N treatment applied to a half-root system may either fully satisfy the plant N demand (N satiety) or result in plant N limitation (N deficit). Plant N satiety resulted in nodule senescence whereas plant N limitation stimulates the expansion of mature nodules. The development of the nodules in untreated roots is strongly dependent on the level of supply of the whole plant, revealing the control by the systemic signaling of the whole plant N demand.

Symbiotic development and function are tightly correlated to carbon allocation from the shoot to the symbiotic organs

Several ¹⁴CO₂ pulse-chase studies have shown that sucrose from photosynthesis is delivered to the nodules via the phloem and then degraded in the cytosol of plant cells to produce organic acids (Gordon et al., 1985; Streeter and Wong, 1988; Rosendahl et al., 1990; Figure 4). Sucrose is first metabolized to UDP-glucose plus fructose, via sucrose synthase (SS), and then oxidized through glycolysis to phosphoenolpyruvate (PEP). PEP is then successively metabolized to oxaloacetate (OAA) and malate by PEP carboxylase (PEPC) and malate dehydrogenase (MDH), respectively. In the different types of nodules, determined or undetermined, the transport of sucrose and/or organic acids to infected cells is preferentially symplastic or apoplastic, or a combination of both (Booth et al., 2021). Thus, the presence of plasmodesmata in soybean (Brown et al., 1995), faba bean (Abd-Alla et al., 2000), and Medicago truncatula (Gaudioso-Pedraza et al., 2018) suggests symplastic transport of C metabolites. Furthermore, the high expression of SWEET-type transporters in vascular parenchyma cells of M. truncatula or Lotus japonicus (Kryvoruchko et al., 2016; Sugiyama et al., 2017) and ALMT (Takanashi et al., 2016) suggests that there is also an apoplastic pathway for the delivery of either sucrose or organic acids to infected cells. Analysis of the expression and activity of enzymes involved in the conversion of sugars to organic acids shows that both vascular parenchyma, non-infected and infected cells are involved in the process of degrading sucrose to malate, albeit in different ways depending on the nodule type (Vance, 2008; Booth et al., 2021). In the determinate nodules of soybean and chickpea, most of the carbon metabolism occurs in non-infected and vascular parenchyma cells (Kouchi et al., 1988; Day and Copeland, 1991), whereas in indeterminate nodules, such as in pea and M. sativa, or in L. japonicus, it is more evenly distributed between infected and non-infected cells and vascular parenchyma cells (Fedorova et al., 1999; Hohnjec et al., 2003; Takanashi et al., 2012). Microoxic conditions associated with O₂ channeling by leghemoglobins prevail in central cells of the nodule as nitrogenase requires a very low O₂ level to be active. In infected cells, some of the malate is transported to the mitochondria where it is used to regenerate ATP and produce carbon skeletons needed for the assimilation of the reduced nitrogen, i.e., NH₄⁺, produced in the bacteroids (Gordon et al., 1985; Rosendahl et al., 1990; Smith et al., 2002). Nodule mitochondria are characterized by their ability to produce ATP more efficiently and at lower O₂ levels than mitochondria in roots and other tissues (Booth et al., 2021). Furthermore, the operation of a phytoglobin-nitric oxide (Pgb-NO) respiration pathway, in which O₂ is consumed by phytoglobins whose affinity for oxygen (K_d O₂ ≈ 2–10 nM) is significantly higher than that of cytochrome oxidase (K_d O₂ ≈ 50–200 nM), allows the innermost cells of nodules to regenerate ATP under the microoxic conditions prevailing in nodules (Berger et al., 2019; Berger et al., 2021). The question particularly arises as to what the O₂ concentration is and what type of respiration is functioning in the infected and uninfected cells in the fixation zone. Indeed, measurements with microelectrodes in determinate nodules such as soybean (Tjepkema and Yocum, 1974) and L. japonicus (Denison, 1992), as well as in indeterminate nodules of M. sativa (Soupène et al., 1995), revealed a strong O₂ gradient between the outside and inside of the nodules. However, to our knowledge, there is no direct way to investigate the difference in O₂ concentration between infected and uninfected cells in vivo. Considering that leghemoglobins are exclusively localized in infected cells (Robertson et al., 1984), an O₂ gradient between infected and uninfected cells could be hypothesized. O₂ concentration in infected cells has been indirectly calculated by the fractional oxygenation of leghemoglobin (Denison and Layzell, 1991). In addition, several modeling studies have reported that pO₂ ranges from 12 to 25 µm in the gas spaces surrounding infected and uninfected cells to 10–60 nM in infected cells (Thumfort et al., 1994; Thumfort et al., 1999; Thumfort et al., 2000). Some studies have indirectly addressed the issue of pO₂ differences between infected and uninfected cells in alfalfa (Arrese-Igor et al., 1993), cowpea (Dakora and Atkins, 1990), or soybean (James et al., 1991) and led to the same conclusion of an O₂ gradient within the fixation zone. Thus, it is very likely that depending on the local O₂ concentration, the involvement of Pgb-NO respiration, alongside classical O₂-dependent respiration, might be important for maintaining the energy state and the metabolism of cells (Berger et al., 2019; Berger et al., 2021). Mitochondria also have high MDH activity and low malic enzyme (ME) activity (Day and Mannix, 1988; Bryce and Day, 1990), which favors the reduction of malate to OAA for subsequent ammonia assimilation (Figure 4). Another part of the organic acids is transported in the bacteroids as dicarboxylate, mainly malate (Figure 4; Booth et al., 2021). The activity of the dicarboxylate transporter on the symbiosome membrane has been demonstrated, but the protein has not yet been identified. In contrast, the bacteroid membrane transporter, DctA, has been well characterized and identified in Rhizobium leguminosarum and Bradyrhizobium japonicum (Ronson et al., 1984; Pessi et al., 2007). DctA is upregulated and accounts for most of the carbon influx into the symbiosomes under symbiotic conditions. Once inside the bacteroid, malate is metabolized by the malic enzyme (ME) and MDH to produce pyruvate and OAA, respectively, which fuel the energy and carbon metabolism of the bacteroids (production of ATP and reducing power, storage of excess carbon in the form of carbon polymers, glycogen, and lipids), and allows the reduction of N₂ to NH₃ by nitrogenase (Lodwig et al., 2005; Terpolilli et al., 2016; Liu et al., 2018). Nitrogenase is the major bacteroid process of ATP and reducing power consumption (16 ATP and 8 e- per N₂ fixed; Figure 4). A comprehensive study, combining experimental and metabolic modeling approaches, was recently conducted in R. leguminosarum and Azorhizobium caulinodans to explain the fundamental features of bacteroid metabolism (Schulte et al., 2021). The catabolism of dicarboxylates provides energy for N reduction and allows the synthesis of carbon polymers and alanine. Metabolic modeling of TCA cycle in the bacteroid suggests that catabolism of dicarboxylates induces a higher NADH/NAD ratio than it might have been if fueled by sugars (Schulte et al., 2021). In this study, the authors show that the entire process is dependent on the O₂ concentration, the low O₂ content of which protects nitrogenase from inhibition, reduces the assimilation of NH₄⁺ into glutamate in the bacteroids, and promotes the export of NH₄⁺ and alanine to the plant cell (Schulte et al., 2021). NH₄⁺ exported by the bacteroid is transported into the cytosol of the infected host cells for its assimilation in amino acids, via the combined action of GS/GOGAT (Figure 4; Cordoba et al., 2003; Seabra et al., 2012). This assimilation process requires an additional flux of C skeleton (α-ketoglutarate) provided by photosynthesis to match the NH₄⁺ flux. Assimilated N is then exported from the nodules to the leaves either as amino acids (mainly asparagine) in indeterminate nodules or as ureides in some determinate nodules (Vance, 2008; Sprent, 2009; Liu et al., 2018). The functioning of the SNF results in a combined action of C and N metabolisms, associated with a strong increase in the expression and activity of their primary enzymes as compared to roots (Vance, 2008), emphasizing the interrelation and co-regulation of the two metabolisms.

Figure 4

The control of symbiosis by plant N demand involves systemic signaling

Mineral nitrogen and particularly NO₃⁻ is known to control its root acquisition at multiple levels. These controls may be related to the ion itself or to the product of its assimilation (Ruffel et al., 2008; Ruffel et al., 2014). These controls are exerted at the level of the organ in presence of the ions and involve local signaling, or at the level of the whole plant and involves systemic signaling. Split-root systems have been used to discriminate between these local and systemic signaling (Figure 3). NO₃⁻ itself stimulates root development and induces root activities involved in its uptake and assimilation. It is a common feature of many NO₃⁻ transporters and enzymes involved in NO₃⁻ assimilation to be activated by the presence of the ion (Vidal et al., 2020). Root NO₃⁻ acquisition induction generally requires the presence of the ion at the site of response (local effect) and may occur in absence of assimilation of the ion (Zhang et al., 1999; Wang et al., 2004). At the whole plant level, the main nitrogen sources (NO₃⁻, NH₄⁺, N₂, or amino acids) are finally assimilated into the same downstream N metabolites to fulfill the N requirement generated by plant growth and functioning. The concept of N demand refers to the balance between the N requirement to fulfill the plant growth potential and plant N acquisition capacity (Imsande and Touraine, 1994; Gojon et al., 2009). Plant N demand may be contrasted according to plant growth rates and/or plant N regimes. Plant N satiety is reached when there is full satisfaction of the N demand (excess of mineral nitrogen for example), whereas plant N deficit occurs when the N demand is not fully satisfied, and the N provision limits plant growth. Variation of the level of downstream N metabolites (namely, amino acids) of the shoots translocated in roots may be associated with this variation of N demand, consistent with the hypothesis of amino acids cycling through the phloem being a signal of plant N demand (Imsande, 1986; Muller and Touraine, 1992; Parsons et al., 1993; Tillard et al., 1998; Girin et al., 2010). The control of NO₃⁻ uptake by a systemic signaling of the plant N demand has been evidenced in several biological systems (Gansel et al., 2001; Girin et al., 2007; Ruffel et al., 2008). N-satiety signaling represses NO₃⁻ transporters, while N-deficit signaling upregulates them. Evidence of regulation of root development by similar systemic control has been also evidenced (Forde, 2002). Although mechanisms behind local and systemic N signaling can be discriminated, they share many targets and, at the whole plant level, generally coexist and interact actively. Without specific experimental designs such as plant cultivated in split-root systems, it is therefore difficult to discriminate between these two modes (Figure 3).

There is little evidence on the impact of the plant N status on the early interactions between rhizobium and legume roots (Grillo et al., 2016). However, following the early interaction, the development of symbiosis requires a whole plant N deficit and is suppressed when plant is supplied by high mineral N supply (Streeter and Wong, 1988; Pervent et al., 2021). However, the plant N deficit must not be too extreme because when the seed reserves are totally consumed, nitrogen and carbon metabolites are still needed to form the new symbiotic structures. This argues for the empiric practice used by legume growers of adding a little amount of mineral nitrogen fertilizer as a “starter” at sowing before symbiosis establishment (Imsande, 1986; Streeter and Wong, 1988). Nevertheless, if seed reserves are consumed and if the plant is not able to fulfill its N demand with the mineral N, active symbiosis may be established. The use of split-root systems demonstrated that the control of nodulation by N demand is mainly exerted at the whole plant level (Figure 3A). Nodule formation requires whole plant N-deficit systemic signaling (Streeter and Wong, 1988; Pervent et al., 2021). Split-root studies in M. truncatula did not argue for a strong local effect of NO₃⁻ itself on nodule formation (Pervent et al., 2021), suggesting that repression of nodulation by NO₃⁻ is mainly related to the downstream N-metabolite production in the whole plant rather than to its presence in the nodule environment. The responses of M. truncatula root to systemic N signaling during the interaction with rhizobium and the nodule formation process were characterized (Pervent et al., 2021). The accumulation of many transcripts associated with the transcriptome reprogramming in response to rhizobium requires systemic signaling of N deficit and is repressed by systemic N signaling of N satiety. However, it is likely that systemic N signaling tunes the progression of the process rather than determine a capacity of the plant to respond to rhizobium (Pervent et al., 2021). Globally, the impact of systemic N signaling is more pronounced during nodule organogenesis, bacteroid differentiation, and activation of nitrogen fixation than on early phases of the interaction.

Evidence for a local regulation of mature symbiotic organs by the efficiency of SNF has been reported. Suppressing nitrogen fixation by Ar/O₂ treatments in split-root systems results locally in a rapid inhibition of nodule growth (Singleton and van Kessel, 1987; Kiers et al., 2003; Jeudy et al., 2010). The general small size and the early developmental arrest of nodules formed by fix− bacteria are globally in agreement with an inhibition of nodule development in the absence of N₂ fixation (Laguerre et al., 2012). It was proposed that the plant develops a local nodule autonomous mechanism to restrict the development of nodules formed with ineffective bacteria (Kiers et al., 2003; Oono et al., 2011). Long-term Ar/O₂ treatments of determinate nodules resulted in a decrease of bacteroid fitness in nodules, associated with early nodule senescence (Kiers et al., 2003; Oono et al., 2009). This result was interpreted as a “host sanction” toward the less beneficial partners. In an evolutionary point of view, “sanction” tends to limit ineffective rhizobia multiplication and dispersion when they are released in the soil. The concept has been extended to indeterminate nodules although bacteroids are terminally differentiated and only undifferentiated bacteria are able to multiply (Oono and Denison, 2010; Oono et al., 2011). Although several reports in soybean/Bradyrhizobium and Medicago/Sinorhizobium symbioses indicate that mutations suppressing N₂ fixation do not necessarily impact the reproductive fitness of rhizobia present in nodule (Marco et al., 2009; Laguerre et al., 2012), evidence for a long-term stimulation of the reproductive fitness of fix⁺ rhizobia versus fix− rhizobia in co-infected root system of Mimosa pudica/Cupriavidus taiwanensis have been also reported (Daubech et al., 2017). Mechanisms related to nodule oxygen permeability or pH have been proposed to be instrumental in the local control of legume–rhizobium symbiotic organs (Hunt and Layzell, 1993; Kiers et al., 2003) but, to date, they remain to be precisely elucidated: whether they are the cause, or the consequence of nitrogen fixation inhibition remains unknown.

Mature nodules are also under the control of systemic signaling of the whole plant N demand (Figures 3, 5, 6). SNF is highly sensitive to abiotic stress that may locally suppress plant N acquisition capacity (Durand et al., 1987; Marino et al., 2013; Gil-Quintana et al., 2013a), resulting in whole plant N deficit (Figure 5). Local suppression of N₂ fixation in split-root systems by Ar/O₂ treatment or by inoculation with fix− bacteria of a fix− half-root system results in a compensatory response on the remaining fix+ half-root system (Figures 3B, 5; Jeudy et al., 2010; Laguerre et al., 2012). The stimulation of mature nodule expansion and the formation of new symbiotic organs are observed (Jeudy et al., 2010; Laguerre et al., 2012), both tending to increase the nitrogen fixation of the fix+ half-root system (Figures 3B, 5). This is of biological importance as plants are sessile organisms; they face soil conditions highly variable in time and space. This systemic mechanism contributes to adjust the root N₂ fixation capacity to the whole plant N demand and to symbiotic root “foraging”. However, it operates probably at the root bundle rather than at the nodule level (Laguerre et al., 2012). Plant facing a uniform reduction of its symbiotic root capacity (fix+ and fix− nodules uniformly distributed on the root system) is unable to trigger the systemic response probably because the plant cannot discriminate between efficient and inefficient root bundles and allocate resources preferentially to the efficient ones (Laguerre et al., 2012). The counterpart of the systemic stimulation of symbiosis by plant N deficit is its systemic repression by plant N satiety (Figures 3, 6). The supply of high level of mineral to a half-root system of N₂ fixing plants cultivated in a split-root system results in systemic N-satiety signaling, represses nitrogen fixation, and activates the senescence of the nodules and the degradation of nitrogen-fixing bacteroids (Figures 3, 6; Pérez Guerra et al., 2010; Lambert et al., 2020). A control of symbiotic activity by downstream N metabolites produced in the shoots and translocated to the roots by the phloem has been frequently suggested (Parsons et al., 1993; Imsande and Touraine, 1994; Bacanamwo and Harper, 1997; Neo and Layzell, 1997). Amino acid supply has indeed a strong inhibitory effect on symbiosis (Bacanamwo and Harper, 1997). However, because amino acids may also be a source of nitrogen metabolized by the plant roots, additional evidence is required to confirm this “feedback” model. Transcriptome analysis of M. truncatula plants cultivated in split-root systems revealed that N-demand systemic signaling is a major driver of nodule development and functioning (Figures 5, 6; Lambert et al., 2020). Although plant N satiety activates a general bacterial transcript breakdown associated with bacteroid lysis during nodule senescence, there is little evidence of a gene-specific regulatory effect of systemic signaling of plant N demand on bacteroid transcriptome. However, plant N satiety and plant N-deficit systemic signaling respectively activates and downregulates numerous plant transcripts involved in nodule senescence, while respectively inhibiting and activating transcript families involved in bacteroid differentiation, nodule meristematic cell division, leghemoglobin synthesis, and sucrose transport (Lambert et al., 2020). They are also associated with large reprogramming of hormonal and plant defense genes (Lambert et al., 2020).

Figure 5

Figure 6

Mechanisms underlying the regulation of symbiosis by photosynthesis

Active nodules require a large flux of sucrose from the plant to fuel the N₂ fixation in the bacteroids (energy and reducing power) and to assimilate the NH₄⁺ released in plant cells. Despite numerous physiological evidence highlighting the tight coordination of photosynthesis and symbiotic activity, mechanisms responsible for the control of symbiosis by the plant C status remain unknown at the genetic and/or molecular levels. Both N-deficit and N-satiety signaling were associated with rapid variations of sucrose allocation from the plant to the active nodule (Lambert et al., 2020). This suggested the hypothesis that plant sucrose allocation is a systemic signal that modulates nodule activity as a function of plant N demand (Figure 7). Consistently with this model, sucrose, produced by shoot photosynthesis, is a major metabolite of phloem sap and its flux is expected to be correlated to the plant growth capacity. Interestingly, nodule Sweet sucrose transporters transcripts have been identified as potential targets of both N-satiety and N-deficit systemic signaling, supporting this hypothesis (Lambert et al., 2020). However, whether sucrose allocation variation is a signal by itself or the consequence of another signaling mechanism remains unknown. The central role of nodule sucrose in the control of symbiosis was already suggested by earlier studies in N-limited supply conditions and in response to drought (Baier et al., 2007). The drop in SNF in response to water stress correlated in several grain legume species (soybean, pea, and bean) to the rapid decline of Susy activity leading to sugar accumulation and organic acid depletion in the nodules (González et al., 1995; González et al., 1998; Gordon et al., 1999; Gálvez et al., 2005). This suggested a key role for Susy in the regulation of SNF by carbon. However, in M. sativa and M. truncatula, the drop in Susy activity only occurs after inhibition of SNF, which questions the possible role of the Susy in this regulation in forage legumes (Naya et al., 2007; Larrainzar et al., 2009).

Figure 7

Lessons may be learned from studies on the mechanisms regulating the acquisition of N by C in non-symbiotic plants (review by Chaput et al., 2020). Sugars from photosynthesis or storage organs are known to be regulators of plant metabolism and plant gene expression (Rolland et al., 2006; Eveland and Jackson, 2012). Sucrose, transported from source tissues to sink organs, is first hydrolyzed either by Susy (to UDP-glucose and fructose) or by invertase (to glucose and fructose) before entering cellular metabolism, to be ultimately oxidized to CO₂ through respiration. Multiple levels of sugar sensing have been characterized: a sucrose sensing pathway via a yet unknown sensor (Vaughn et al., 2002); the hexokinase pathway, which, independently of its glucose phosphorylation activity, has glucose sensing activity (Jang et al., 1997; Granot et al., 2013); a hexokinase-independent pathway, probably linked to the regulator of G-protein signaling (RGS1) located on the plasma membrane (Grigston et al., 2008; Urano et al., 2012); a glycolysis-dependent pathway downstream of hexokinase (Xiao et al., 2000); a trehalose pathway (Lunn et al., 2014); and a pathway related to the supply of carbon substrates to mitochondrial respiration (Aubert et al., 1996). As regards N acquisition, several investigations carried out in Arabidopsis roots have shown that the expression of the NO₃⁻ transporter genes NRT2.1 and NPF6.3 was directly related to the concentration of glucose-6-phosphate in the roots (Lejay et al., 2003; Lejay et al., 2008). The use of 6-aminonicotinamide, an inhibitor of glucose-6-phosphate dehydrogenase (G6PDH) and 6-phosphogluconate dehydrogenase (6PGDH), two enzymes of the oxidative part of the pentose phosphate pathway (OPPP), as well as the use of a knockdown mutant for plastid 6-phosphogluconolactonase (PGL3), respectively made it possible to highlight the role of OPPP in the regulation of the expression of NRT2.1 and NPF6.3 (Lejay et al., 2008), as well as nitrate reductase (NR) and nitrite reductase (NiR; Bussell et al., 2013). Together with several studies highlighting the role of OPPP and sugars in the regulation of transporters involved in N acquisition (Oji et al., 1985; Bowsher et al., 1989; Bowsher et al., 1992; Neuhaus and Emes, 2000), these investigations supported the existence of an OPPP-dependent sugar signaling pathway for the regulation of plant N acquisition by roots. In plants, OPPP is the main NADPH regeneration pathway that helps maintain cellular redox balance, especially under oxidative stress. An increased flux through the OPPP results in an increased NADPH/NADP ratio and a better resistance to oxidative stress (Ralser et al., 2007), whereas the mutation of G6PDH, which determines the level of NADPH by controlling the flux of G6P that enters the OPPP, leads to a lower resistance to stress (Juhnke et al., 1996). In Arabidopsis, the recent demonstration of the regulation of AtNRT2.1 by the redox status (Bellegarde et al., 2019) suggests that OPPP, via the regeneration of NADPH, could be the intermediary in C signaling. The chromatin factor HIGH NITROGEN INSENSITIVE9 (HNI9), encoded by a genetically identified regulatory locus of AtNRT2.1, was found to reduce the ROS levels under high, but not low, N provision (Widiez et al., 2011; Bellegarde et al., 2019). Interestingly, in Arabidopsis, the bZIP transcription factor ELONGATED HYPOCOTYL5 (HY5) was shown to be a shoot-to-root mobile systemic signal that mediates light promotion of root growth and NO₃⁻ uptake via the activation of NRT2.1 (Chen et al., 2016). In the shoot, HY5 promotes indirectly carbon assimilation and translocation, whereas in the root, HY5-dependent upregulation of NRT2.1 and NO₃⁻ uptake are favored by an increase in photosynthesis-derived sugars. Together with HNI9, HY5 is required for activation of the detoxification ROS program under high N (Bellegarde et al., 2019). The ability of HY5 to bind the promoter G-box of ROS-responsive genes and regulate de-etiolation in response to light and ROS suggests that HY5 could be involved in the crosstalk between sugars and redox state for the regulation of NRT2.1 and several other NO₃⁻ transporter genes by C through the OPPP (Chen et al., 2016; Gangappa and Botto, 2016; Chaput et al., 2020). This knowledge acquired in Arabidopsis on the regulation of NO₃⁻ acquisition provides a basis to propose a hypothetical model for the regulation of symbiosis by photosynthesis (Figure 8). Interestingly, in soybean, the HY5 ortholog light-induced TGACG-motif binding factor 3/4 (GmSTF3/4) and FLOWERING LOCUS T (GmFTs) were shown to interdependently induce nodule organogenesis (Wang et al., 2021), supporting the idea that these transcription factors could also be part of the systemic regulation of symbiosis by C (Figure 8). OPPP also has a major role in symbiosis (Figure 8). NADPH is the primary redox cofactor that regulates the regeneration of glutathione and reduced ascorbate, which, in turn, act as secondary redox cofactors in the turnover, or even the detoxification, of reactive oxygen species (ROS) and reactive nitrogen species (RNS) (Noctor and Foyer, 1998; Apel and Hirt, 2004). Interestingly, ROS and nitric oxide (NO), as well as glutathione and homoglutathione, have been shown to be major regulators of symbiosis establishment and functioning (Pauly et al., 2006; Puppo et al., 2013; Berger et al., 2019). In mature nodules, NADPH oxidases (RBOHs) are major sources of H₂O₂ production, via superoxide anion dismutation (Marino et al., 2011; Arthikala et al., 2014). NR and electron transfer chains from both plant and bacterial partners significantly contribute to NO production in N₂-fixing nodules (Sánchez et al., 2010; Horchani et al., 2011; Berger et al., 2021). The NO concentration is itself finely regulated by the phytoglobin Pgb1.1, whose function is to allow NO to exercise its signaling and metabolic intermediary functions at the different stages of the symbiosis without reaching toxic levels for the metabolism (Fukudome et al., 2016; Fukudome et al., 2019; Berger et al., 2020). In this reaction, NO is first oxidized to NO₃⁻ by oxyPgb, which is converted to metPgb. MetPgb is then reduced by a MetPgb reductase (MetPgb-R) at the expense of NAD(P)H reducing power (Igamberdiev et al., 2006). Both S-sulfenylated and S-nitrosylated proteins, resulting from post-translational modifications generated by H₂O₂ and NO, have been detected during early interaction and in functioning nodules, linking ROS/NO production to redox-based protein regulation (Puppo et al., 2013). Thus, considered together, many studies allow to make functional links between carbon metabolism, the regeneration of NADPH and the regulation of redox status by OPPP, and the regulation of SNF by ROS and NO in mature nodules (Figure 8). However, this model remains highly speculative and demonstration of such mechanisms remains to be done.

Figure 8

Multiple pathways are involved in the systemic control of symbiosis

In the last decade, significant discoveries allowed the characterization of receptors, peptides, and transduction pathways involved in the systemic control of nodule formation (Figure 9). Nevertheless, how these mechanisms are integrated at the whole plant level and contribute to the global phenotypes in response to variation of plant N demand and photosynthesis remains elusive. AON was shown to result in the inhibition of nodule formation by a pre-existing nodule (Kosslak et al., 1983; Kosslak and Bohlool, 1984; Mathews et al., 1989; Olsson et al., 1989; Kassaw et al., 2015). Evidenced by split-root experiments, this regulation involves systemic signaling between shoots and roots (Kosslak and Bohlool, 1984; Olsson et al., 1989; Kassaw et al., 2015). Pioneer genetic studies in several legume species allowed the identification of AON components (Caetano-Anollés and Gresshoff, 1991; Sagan et al., 1995; Wopereis et al., 2000; Chaulagain and Frugoli, 2021). AON mutants form generally more nodules than wild type and therefore display super/hyper nodulation phenotypes (Caetano-Anollés and Gresshoff, 1991; Sagan et al., 1995). Several recent reviews described in detail our current knowledge of the related molecular mechanisms (Figure 9; Chaulagain and Frugoli, 2021; Gautrat et al., 2021; Roy and Müller, 2022). AON involves CLV3-like 12-amino acid peptides (CLE) synthetized in roots, translocated by the xylem flux to the shoots, where they bind CLV3-like Leucine Rich Repeat Receptors Like Kinases LRR-RLK (Figure 9). SUNN, HAR1, and NARK loci encode these AON LRR-RLK in M. truncatula, L. japonicus, and soybean, respectively (Krusell et al., 2002; Nishimura et al., 2002; Searle et al., 2003; Schnabel et al., 2005). The receptors exist as homodimers or heterodimers formed with truncated co-receptors (LjCLV2, LjKLV in L. japonicus or MtCLV2 and MtCRN in M. truncatula; Miyazawa et al., 2010; Krusell et al., 2011; Crook et al., 2016). CLE peptides are encoded by large gene families in legumes and non-legume plants (Yamaguchi et al., 2016). The role of CLE peptides in the control of nodule number was demonstrated only for a few of them: MtCLE12, MtCLE13, and MtCLE35 in M. truncatula (Mortier et al., 2010; Mortier et al., 2012; Mens et al., 2020); LjCLE-RS1, LjCLE-RS2, and LjCLE-RS3 in L. japonicus (Okamoto et al., 2009; Nishida et al., 2016); and Gm-RIC1 and GmRIC2 in soybean (Reid et al., 2011a). The corresponding genes are upregulated in the roots in response to the rhizobium/nodule formation (Figure 9). The interaction of AON LRR-RLK receptor and CLE peptides activates the shoot-derived systemic inhibition of nodulation (Figure 8). A downstream component of AON LRR-RLK receptors is the small RNA miR2111 (Figure 9; Tsikou et al., 2018). miR2111 is synthetized as a precursor in the shoots, processed and translocated to the roots by the phloem. In the root, miR2111 post-transcriptionally represses TML1 and 2 genes, encoding ubiquitin ligases, resulting in the inhibition of nodule formation according to a still unknown mechanism (Figure 9; Magori et al., 2009; Takahara et al., 2013; Tsikou et al., 2018; Gautrat et al., 2019). Other signaling processes may also be implicated downstream of AON LRR-RLK receptors. In L. japonicus, the symbiosis establishment results in a HAR1-dependent upregulation of cytokinin synthesis in the shoots that is implicated in the regulation of nodule formation in roots (Sasaki et al., 2014). In addition, AON was associated in M. truncatula with a reduction of the shoot-to-root transport of auxin (van Noorden et al., 2006). Early studies revealed that AON mutants maintain the ability to form nodules under high NO₃⁻ supply, demonstrating the role of AON in the control of nodulation by the plant N status (Carroll et al., 1985; Sagan et al., 1995). The resistance of nodulation to NO₃⁻ in the sunn mutant of M. truncatula was related to a release of the N-satiety systemic repression (Jeudy et al., 2010). The response of sunn and wild-type roots to systemic N signaling during nodule formation was compared using split-root systems. A role of AON in the control of nodule formation by plant N demand was confirmed but AON-independent components were evidenced (Kassaw et al., 2015; Pervent et al., 2021). The role of AON in the regulation of nodulation by NO₃⁻ was also discussed in L. japonicus (Nishida et al., 2020). Some CLE genes encoding peptides were found to be upregulated by NO₃⁻ (MtCLE35, LjCLE-RS2, LjCLE-RS3, and Gm-NIC1) and to inhibit partial nodulation through the AON LRR-RLK receptor/miR2111/TML pathway (Figure 9; Okamoto et al., 2009; Nishida et al., 2016; Lebedeva et al., 2020; Mens et al., 2020; Moreau et al., 2021). In M. truncatula, only MtTML2 is downregulated in response to MtCLE35 overexpression, suggesting a specificity of the response to NO₃⁻ as compared to the response to rhizobium/nodule formation (Moreau et al., 2021). For decades, AON was the unique identified systemic pathway controlling symbiotic development. Discovery of M. truncatula TR185/cra2 mutants shaded the light on an additional pathway responsible for systemic activation of the root nodulation capacity (Bourion et al., 2014; Huault et al., 2014; Laffont et al., 2019). The mutants display highly branched root phenotype and modified responses to NO₃⁻ in non-symbiotic conditions (Bourion et al., 2014; Huault et al., 2014). Their capacity to form nodules with rhizobium is dramatically impaired due to the absence of a systemic signaling originated from the shoot (Huault et al., 2014; Laffont et al., 2019). The MtCRA2 gene, impaired in the mutants, encodes a Leucine-Rich Repeat Receptor-Like Kinase (LRR-RLK) present in shoots able to interact with peptides of the CEP family (C terminally encoded peptides; Figure 9). Both MtCEP1 and MtCEP7 peptides were found to activate the CRA2 systemic signaling, allowing nodulation (Laffont et al., 2019; Laffont et al., 2020). The two corresponding genes are upregulated in the root in response to the absence of mineral nitrogen and to rhizobium, suggesting a control of the pathway by both nitrogen status of the plant and infection by the bacteria. Intriguingly, MtCEP/MtCRA2 and the AON MtCLE/MtSUNN pathways share the downstream miR2111/TML component but act antagonistically on it (Figure 9). MtCEP/MtCRA2 stimulates the accumulation of miR2111 in the shoot to promote the cleavage MtTML transcript in the root, resulting in a stimulation of the root nodulation capacity (Gautrat et al., 2019; Gautrat et al., 2020). Nevertheless, the relative contribution of MtCEP/MtCRA2, MtCLE/MtSUNN, and possibly other unknown components in the control of nodulation by the N status of the plant is not well understood. Furthermore, the role, if any, of MtCEP/MtCRA2 on the response mature nodules to N demand and mineral N is not known because the mutant is impaired in nodule formation. Physiological and molecular characterization of TR185/cra2 plants in non-symbiotic conditions described a N-limitation phenotype and the effect of the mutation on plant NO₃⁻ acquisition and on root NO₃⁻ transporters’ gene expression. AtCEPR1/AtCEPR2 orthologs of MtCRA2 as well as CEP peptides were identified in Arabidopsis (Tabata et al., 2014). A root target of systemic action of AtCEP1/CEPR1 is the high-affinity NO₃⁻ transporter gene AtNRT2.1, known to be upregulated in response to the whole plant N deficiency. AtCEPD1 and AtCEPD2 (CEP downstream 1 and 2), two putative Class 3 glutaredoxins, might play the role of a systemic phloem signal from shoot to root to upregulate AtNRT2.1 (Ohkubo et al., 2017). High-affinity NO₃⁻ transporters are not the only targets of CEP/CEPR1 and CEP/CRA2 pathways both in Arabidopsis and in non-symbiotic M. truncatula plants, and root architecture was found to be strongly impaired in related mutants (Bourion et al., 2014; Delay et al., 2019; Chapman et al., 2020). How similar are the CEP receptors’ transduction pathways involved in the control of NO₃⁻ uptake and root development in non-symbiotic conditions and the control of nodule formation in symbiotic conditions is not clearly understood. The role of CEPD proteins in the systemic regulation of nodulation downstream of CRA2 in M. truncatula was questioned (Gautrat et al., 2020). Although it cannot be excluded that CRA2 might be a component of the same mechanism adjusting either nodule formation, NO₃⁻ acquisition, or root development to plant N demand, this remains to be demonstrated. Because the inactivation of CEP receptor genes has pleiotropic impacts on plant development and functioning, as well as mineral nitrogen acquisition, discriminating between direct and indirect impacts on the nodule formation phenotype in legumes is difficult. Nevertheless, the dramatic inhibition of nodulation observed in the cra2 mutant cannot be simply explained as the result of the N-deficiency phenotype because plant N deficit stimulates rather than inhibits nodulation. To our knowledge, there is little convincing evidence for an active role of bacteria in the regulation of nodules by the holobiont N status (Lambert et al., 2020). However, several reports indicate that GlnD and PII, two bacterial regulatory components controlling nitrogen metabolism in bacteria, are required for symbiosis functioning, suggesting that the question might deserve more investigation (Arcondéguy et al., 1997; Yurgel et al., 2012; D’Apuzzo et al., 2015).

Figure 9

Does the local sensing of NO₃⁻ contribute to the control of symbiosis by plant N demand?

In addition to its role as a resource for downstream N-metabolite synthesis, NO₃⁻ itself plays a role of signaling molecule in plant organs (review by Maghiaoui et al., 2020). In Arabidopsis, the use of null NR mutants’ NO₃⁻-specific effects is independent of its reduction (Zhang et al., 1999; Wang et al., 2004). The complex mechanisms related to root NO₃⁻ sensing begin to unravel in Arabidopsis (Maghiaoui et al., 2020). The NO₃⁻ transporter AtNRT1.1 (NPF6.3/CHL1) plays the role of sensor and governs a wide range of response to NO₃⁻ independently of its transport activity and NO₃⁻ assimilation (Ho et al., 2009; Krouk et al., 2010; Bouguyon et al., 2015; Riveras et al., 2015; Maghiaoui et al., 2020). Other central players are some NLP transcription factors required for the induction of many target genes by NO₃⁻ including those responsible for its reduction and assimilation (Castaings et al., 2009; Marchive et al., 2013; Guan et al., 2017; Liu et al., 2017; Liu et al., 2022). Several studies described the regulation in legumes of both nodule formation and functioning by NO₃⁻ through the action of NLP proteins (Figure 9; Lin et al., 2018; Nishida et al., 2018; Moreau et al., 2021). In M. truncatula, MtNLP1, activated in response to NO₃⁻, and MtNIN, required for nodulation in the presence of rhizobium, were shown to antagonistically interact for the transcriptional activation of key genes (Figure 9; Lin et al., 2018). NLP proteins (MtNLP1 and LjNRSYM1) were implicated in the transcriptional activation of NO₃⁻-responsive genes, including MtCLE35 and LjCLE-RS2, as well as NR and NiR genes (Nishida et al., 2018; Moreau et al., 2021). This local activation of CLE genes was shown to be associated with activating the LjHAR1/MtSUNN AON LRR-RLK-dependent systemic inhibition of nodulation in model legumes (Figure 9). Nevertheless, the biological significance of the hypothesis of systemic inhibition of nodulation activated locally by NO₃⁻ remains elusive because (1) split-root studies rather suggest a regulation of nodulation by N demand related to downstream N-metabolite production at the whole plant level (Pervent et al., 2021) and (2) such mechanism does not explain inhibition of symbiosis by other N sources such as amino acids or NH₄⁺ (Yamashita et al., 2019). Furthermore, because of their roles in the activation of NO₃⁻ assimilation, an indirect impact of NLPs on downstream N-metabolite synthesis cannot be ruled out. The use of legume mutant background impaired in NO₃⁻ reduction such as null NR mutants may unequivocally discriminate between a role of these NLPs in the inhibition of nodulation by NO₃⁻ itself or by products of its assimilation. Finally, there are intriguing reports showing that, in some conditions, NO₃⁻ may be required for optimal nitrogen fixation in mature nodules of L. japonicus through the action of specific nodule NO₃⁻ transporters (Valkov et al., 2017; Valkov et al., 2020). This raised the hypothesis of a control of symbiotic activity by the NO₃⁻ flux fueling the Pgb-NO respiration known to be active in the microoxic conditions of mature nodules (Horchani et al., 2011).

Perspectives

The last decade yielded important knowledge on multiple mechanisms involved in the adjustment of the symbiotic capacity to the plant nutritional demand as a function of the plant environment. Because symbiosis allows the plant to acquire N from air at the expense of photosynthates, the plant N and C status are major drivers of these mechanisms. Local environmental conditions are tuning the adjustment of symbiosis activity to the whole plant nutritional status not only through the availability of N and C resources (mineral nitrogen, light, and CO₂) but also by allowing or inhibiting the development and/or functioning of symbiotic organs. Plants continuously adapt to these conditions that are frequently heterogeneous in space and time. The nutritional demand is therefore necessarily integrated at the level of the whole plant, resulting in foraging responses either by stimulating symbiotic capacity (under N-deficit or eCO₂ conditions) or by inhibiting it (under N-satiety or low-light conditions). These responses are activated by both systemic and local signaling pathways. The discovery of multiple pathways, acting simultaneously and targeting almost all aspects of nodule development and functioning, revealed not only the central role of the adjustment of the symbiotic capacity to the plant nutritional demand, but also its extraordinary complexity. However, the biological impact of these pathways and their relative role in the whole plant phenotype as a function of the environment is far to be understood.

Up to now, most investigations mainly focused on regulatory circuits controlling early plant–rhizobium interaction and nodule formation. Because the nodule formation process is associated with the activation of a large set of specific genes, earlier studies have predicted that specific symbiotic mechanisms may be operating in this control (Ruffel et al., 2008). However, the current knowledge prompted us to modulate this interpretation. Although the MtCEP/MtCRA2, the AON MtCLE/MtSUNN, or the NLP-related NO₃⁻-responsive pathways target many specific symbiotic genes and development processes in legume plants, there is increasing evidence indicating that (1) legume mutants impaired in these pathways display non-symbiotic phenotypes often related to N nutrition, and (2) these pathways belong to families of pathways present in non-legumes regulating root development and mineral nitrogen acquisition in response to NO₃⁻. The HY5 pathway was found to be important not only for the systemic regulation by light of NO₃⁻ acquisition under non-symbiotic conditions but also for nodule formation under symbiotic conditions. A major challenge for future studies will be to revisit the plant phenotypes and discriminate between the pleiotropic consequences of mutations impairing the functions of these pathways. How much the impact of the Mtcra2 mutation on non-symbiotic functions (nitrogen limitation and root development) might influence the nodule formation phenotype of the mutant remains an open question. We do not know if, in NLP mutants (LjNRsym1 and Mtnlp1), the reduced activation by NO₃⁻ of the NO₃⁻ assimilation pathway, which is expected to lower the levels of downstream N metabolites, contributes to reduce the response of nodulation to NO₃⁻. Discriminating between the effects of NO₃⁻ itself and its assimilation on the regulation of nodulation will require appropriate strategies (NR mutants, for example). Although many progresses have been made in our understanding of AON, several studies suggest that some pieces of the puzzle in the control nodule formation by systemic signaling N demand remain unknown, particularly the mechanisms involved in the bacteroid differentiation and in the activation of nitrogen fixation in newly formed nodules (Pervent et al., 2021). Furthermore, although the last decade yields the discoveries of CLE and CEP peptides as well as miR2111, playing the role of signal molecules between root and shoot, the role of the other plant hormones in the systemic control of nodule formation by plant N demand remains to be clarified. Although ethylene has been implicated in the control of nodulation (particularly infection), its role in the plant response to N and C status through systemic signaling regulation deserves further investigation (Penmetsa et al., 2003; Prayitno et al., 2006; Zhu et al., 2020).

Less attention was made on the control of mature nodule development and functioning by the whole plant nutritional status. However, the N and C status of the plant may strongly determine mature nodule behavior, either by stimulating nodule expansion or by activating nodule senescence. Consistently with the tight integration of N and C signaling, the regulation of sucrose allocation to the nodule was associated with N-satiety and N-deficit systemic signaling, suggesting that the fueling symbiosis by C metabolites may contribute to a systemic N-demand signaling process. (Jeudy et al., 2010; Lambert et al., 2020). Supporting this hypothesis, transcripts encoding sucrose transporters likely responsible for the nodule acquisition of sucrose are targets of the N-demand systemic signaling pathway (Lambert et al., 2020). However, further studies are required to validate this model and its biological relevance. More globally, there is a lack of knowledge on mechanisms responsible for the coordination of symbiotic activity and photosynthesis. The last decade yielded the discovery of the role of the HY5 pathway in the control of nodule formation (Chen et al., 2016; Wang et al., 2021). However, questions related to biological significance and physiological impact of this pathway in the control of symbiosis deserve further investigations. The HY5 pathway was identified as a response to light, whereas most of the physiological data suggest a control of symbiosis by photosynthates allocated from the shoots to the roots. Furthermore, eCO₂ stimulates symbiosis without any change in light and, therefore, independently of light-induced regulation of HY5. Up to now, most of the reported functions of HY5 relate to nodule formation and little is known about its role in mature nodule functioning. Whether additional function of HY5, or other mechanisms, is involved in the regulation of the symbiosis by photosynthesis and sucrose allocation by the plant remains to be further investigated. The discovery of the role of OPPP and the redox status in the regulation of plant NO₃⁻ acquisition by photosynthesis suggests evaluating its role in the control of symbiosis in legumes. Although the critical role of the redox status (ROS and NOS) in symbiotic development and functioning has been clearly demonstrated, the possible role of the C metabolites’ allocation and the metabolites’ flux through the OPPP (providing reducing power necessary for the control of redox status) is an attracting hypothesis in the context of the regulation of symbiosis by the whole plant nutritional status. One of the major lessons of the last decade is our need to analyze integrated phenotypes of mutants impaired in regulatory components of symbiosis taking into account not only specific symbiotic function but also their possible interactions with whole holobiont development, and considering C and N economy as well as their impact on metabolic fluxes. Several efforts in this direction have been made: (1) modeling C and N exchanges as well as metabolic flux in the context of the symbiotic plant (Salon et al., 2009; Moreau et al., 2012; diCenzo et al., 2016; Schulte et al., 2021) and (2) investigating these regulations using split-root systems, which allows discrimination between local and inter-organ signaling (Ruffel et al., 2008; Jeudy et al., 2010; Laguerre et al., 2012; Lambert et al., 2020; Pervent et al., 2021).

A major biological role of these mechanisms relates to the whole symbiotic plant adaptation to local conditions impairing symbiotic activity (such as drought, salt, temperature, heavy metals, and flooding). As a sessile organism, the holobiont adapts to local stress and circumvents the plant N deficit resulting from local inhibition of SNF by stimulating nodule development in the root, allowing efficient symbiosis (Jeudy et al., 2010; Laguerre et al., 2012). This adaptation to satisfy the N demand is made by allocating preferentially C resources to more efficient roots at the expense of inefficient ones. In the context of climate change, soil conditions will be more heterogeneous and fluctuating than ever, resulting more frequently in local stresses (Dusenge et al., 2019). In addition, atmospheric ambient CO₂ will increase, potentially modifying the conditions of plant C acquisition. A major challenge for plant science is to design strategies to select genotypes better adapted to these new conditions. The potential of legumes in this context has been highlighted because, unlike cereals acquiring mainly NO₃⁻ as a N source, symbiotic N₂-fixing legumes can benefit from elevated CO₂ (Rogers et al., 2009). Despite the major threat of climate change in agriculture, changes in the equilibrium of C/N trade-offs in symbiotic crop legumes may be an opportunity for the selection of new genotypes able to better adapt to soil local constraints and display better competitivity as compared to other C3 non-symbiotic plants.

References

• 1

  Abd-AllaM. H.KoyroH.-W.YanF.SchubertS.PeiterE. (2000). Functional structure of the indeterminate vicia faba l. root nodule: Implications for metabolite transport. J. Plant Physiol.157, 335–343. doi: 10.1016/S0176-1617(00)80056-5

  • CrossRef
  • Google Scholar
  • View reference in article

• 2

  ÅgrenG. I.WetterstedtJ.Å.M.BillbergerM. F. K. (2012). Nutrient limitation on terrestrial plant growth – modeling the interaction between nitrogen and phosphorus. New Phytol.194, 953–960. doi: 10.1111/j.1469-8137.2012.04116.x

  • CrossRef
  • Google Scholar
  • View reference in article

• 3

  ApelK.HirtH. (2004). Reactive oxygen species: Metabolism, oxidative stress, and signal transduction. Annu. Rev. Plant Biol.55, 373–399. doi: 10.1146/annurev.arplant.55.031903.141701

  • CrossRef
  • Google Scholar
  • View reference in article

• 4

  ApplebyC. A. (1992) The origin and functions of haemoglobin in plants. Sci. Progress Oxford (Sage Publications, Ltd.) 76, 365–398. Available at: https://www.jstor.org/stable/43421309.

  • Google Scholar
  • View reference in article

• 5

  ArcondéguyT.HuezI.TillardP.GangneuxC.de BillyF.GojonA.et al. (1997). The rhizobium meliloti PII protein, which controls bacterial nitrogen metabolism, affects alfalfa nodule development. Genes Dev.11, 1194–1206. doi: 10.1101/gad.11.9.1194

  • CrossRef
  • Google Scholar
  • View reference in article

• 6

  Arrese-IgorC.RoyuelaM.de LorenzoC.de FelipeM. R.Aparicio-TejoP. M. (1993). Effect of low rhizosphere oxygen on growth, nitrogen fixation and nodule morphology in lucerne. Physiologia Plantarum89, 55–63. doi: 10.1111/j.1399-3054.1993.tb01786.x

  • CrossRef
  • Google Scholar
  • View reference in article

• 7

  ArthikalaM.-K.Sánchez-LópezR.NavaN.SantanaO.CárdenasL.QuintoC. (2014). RbohB, a phaseolus vulgaris NADPH oxidase gene, enhances symbiosome number, bacteroid size, and nitrogen fixation in nodules and impairs mycorrhizal colonization. New Phytol.202, 886–900. doi: 10.1111/nph.12714

  • CrossRef
  • Google Scholar
  • View reference in article

• 8

  AubertS.GoutE.BlignyR.Marty-MazarsD.BarrieuF.AlabouvetteJ.et al. (1996). Ultrastructural and biochemical characterization of autophagy in higher plant cells subjected to carbon deprivation: Control by the supply of mitochondria with respiratory substrates. J. Cell Biol.133, 1251–1263. doi: 10.1083/jcb.133.6.1251

  • CrossRef
  • Google Scholar
  • View reference in article

• 9

  BacanamwoM.HarperJ. E. (1997). The feedback mechanism of NO₃^- inhibition of nitrogenase activity in soybean may involve asparagine and/or products of its metabolism. Physiologia Plantarum100, 371–377. doi: 10.1111/j.1399-3054.1997.tb04795.x

  • CrossRef
  • Google Scholar
  • View reference in article

• 10

  BaierM. C.BarschA.KüsterH.HohnjecN. (2007). Antisense repression of the medicago truncatula nodule-enhanced sucrose synthase leads to a handicapped nitrogen fixation mirrored by specific alterations in the symbiotic transcriptome and metabolome. Plant Physiol.145, 1600–1618. doi: 10.1104/pp.107.106955

  • CrossRef
  • Google Scholar
  • View reference in article

• 11

  BellegardeF.MaghiaouiA.BoucherezJ.KroukG.LejayL.BachL.et al. (2019). The chromatin factor HNI9 and ELONGATED HYPOCOTYL5 maintain ROS homeostasis under high nitrogen provision. Plant Physiol.180, 582–592. doi: 10.1104/pp.18.01473

  • CrossRef
  • Google Scholar
  • View reference in article

• 12

  BergerA.BoscariA.FrendoP.BrouquisseR. (2019). Nitric oxide signaling, metabolism and toxicity in nitrogen-fixing symbiosis. J. Exp. Bot.70, 4505–4520. doi: 10.1093/jxb/erz159

  • CrossRef
  • Google Scholar
  • View reference in article

• 13

  BergerA.BoscariA.PuppoA.BrouquisseR. (2021). NO3- reductases and hemoglobins control nitrogen-fixing symbiosis by regulating nitric oxide accumulation. J. Exp. Bot.72, 873–884. doi: 10.1093/jxb/eraa403

  • CrossRef
  • Google Scholar
  • View reference in article

• 14

  BergerA.GuinandS.BoscariA.PuppoA.BrouquisseR. (2020). Medicago truncatula phytoglobin 1.1 controls symbiotic nodulation and nitrogen fixation via the regulation of nitric oxide concentration. New Phytol.227, 84–98. doi: 10.1111/nph.16462

  • CrossRef
  • Google Scholar
  • View reference in article

• 15

  BoivinS.Ait LahmidiN.SherlockD.BonhommeM.DijonD.Heulin-GottyK.et al. (2020). Host-specific competitiveness to form nodules in rhizobium leguminosarum symbiovar viciae. New Phytol.226, 555–568. doi: 10.1111/nph.16392

  • CrossRef
  • Google Scholar
  • View reference in article

• 16

  BoothN. J.SmithP. M. C.RameshS. A.DayD. A. (2021). Malate transport and metabolism in nitrogen-fixing legume nodules. Molecules26, 6876. doi: 10.3390/molecules26226876

  • CrossRef
  • Google Scholar
  • View reference in article

• 17

  BouguyonE.BrunF.MeynardD.KubešM.PerventM.LeranS.et al. (2015). Multiple mechanisms of NO3- sensing by arabidopsis NO3- transceptor NRT1.1. Nat. Plants1, 15015. doi: 10.1038/nplants.2015.15

  • CrossRef
  • Google Scholar
  • View reference in article

• 18

  BourionV.Heulin-GottyK.AubertV.TisseyreP.Chabert-MartinelloM.PerventM.et al. (2017). Co-Inoculation of a pea core-collection with diverse rhizobial strains shows competitiveness for nodulation and efficiency of nitrogen fixation are distinct traits in the interaction. Front. Plant Sci.8. doi: 10.3389/fpls.2017.02249

  • CrossRef
  • Google Scholar
  • View reference in article

• 19

  BourionV.MartinC.de LarambergueH.JacquinF.AubertG.Martin-MagnietteM.-L.et al. (2014). Unexpectedly low nitrogen acquisition and absence of root architecture adaptation to NO3- supply in a medicago truncatula highly branched root mutant. J. Exp. Bot.65, 2365–2380. doi: 10.1093/jxb/eru124

  • CrossRef
  • Google Scholar
  • View reference in article

• 20

  BowsherC. G.BoultonE. L.RoseJ.NayagamS.EmesM. J. (1992). Reductant for glutamate synthase in generated by the oxidative pentose phosphate pathway in non-photosynthetic root plastids. Plant J.2, 893–898. doi: 10.1111/j.1365-313X.1992.00893.x

  • CrossRef
  • Google Scholar
  • View reference in article

• 21

  BowsherC. G.HucklesbyD. P.EmesM. J. (1989). Nitrite reduction and carbohydrate metabolism in plastids purified from roots of pisum sativum l. Planta177, 359–366. doi: 10.1007/BF00403594

  • CrossRef
  • Google Scholar
  • View reference in article

• 22

  BrownS. M.OparkaK. J.SprentJ. I.WalshK. B. (1995). Symplastic transport in soybean root nodules. Soil Biol. Biochem.27, 387–399. doi: 10.1016/0038-0717(95)98609-R

  • CrossRef
  • Google Scholar
  • View reference in article

• 23

  BryceJ. H.DayD. A. (1990). Tricarboxylic acid cycle activity in mitochondria from soybean nodules and cotyledons. J. Exp. Bot.41, 961–967. doi: 10.1093/jxb/41.8.961

  • CrossRef
  • Google Scholar
  • View reference in article

• 24

  BussellJ. D.KeechO.FenskeR.SmithS. M. (2013). Requirement for the plastidial oxidative pentose phosphate pathway for NO₃^- assimilation in arabidopsis. Plant J.75, 578–591. doi: 10.1111/tpj.12222

  • CrossRef
  • Google Scholar
  • View reference in article

• 25

  Caetano-AnollésG.GresshoffP. M. (1991). Plant genetic control of nodulation. Annu. Rev. Microbiol.45, 345–382. doi: 10.1146/annurev.mi.45.100191.002021

  • CrossRef
  • Google Scholar
  • View reference in article

• 26

  Caetano-AnollésG.JoshiP. A.GresshoffP. M. (1991). Spontaneous nodules induce feedback suppression of nodulation in alfalfa. Planta183, 77–82. doi: 10.1007/BF00197570

  • CrossRef
  • Google Scholar
  • View reference in article

• 27

  Caetano-AnollésG.LagaresA.BauerW. D. (1990). Rhizobium meliloti exopolysaccharide mutants elicit feedback regulation of nodule formation in alfalfa. Plant Physiol.92, 368–374. doi: 10.1104/pp.92.2.368

  • CrossRef
  • Google Scholar
  • View reference in article

• 28

  CarrollB. J.McNeilD. L.GresshoffP. M. (1985). A supernodulation and NO₃^–Tolerant symbiotic (nts) soybean mutant. Plant Physiol.78, 34–40. doi: 10.1104/pp.78.1.34

  • CrossRef
  • Google Scholar
  • View reference in article

• 29

  CastaingsL.CamargoA.PocholleD.GaudonV.TexierY.Boutet-MerceyS.et al. (2009). The nodule inception-like protein 7 modulates NO₃^- sensing and metabolism in arabidopsis. Plant J.57, 426–435. doi: 10.1111/j.1365-313X.2008.03695.x

  • CrossRef
  • Google Scholar
  • View reference in article

• 30

  ChapmanK.IvanoviciA.TaleskiM.SturrockC. J.NgJ. L. P.Mohd-RadzmanN. A.et al. (2020). CEP receptor signalling controls root system architecture in arabidopsis and medicago. New Phytol.226, 1809–1821. doi: 10.1111/nph.16483

  • CrossRef
  • Google Scholar
  • View reference in article

• 31

  ChaputV.MartinA.LejayL. (2020). Redox metabolism: the hidden player in carbon and nitrogen signaling? J. Exp. Bot.71, 3816–3826. doi: 10.1093/jxb/eraa078

  • CrossRef
  • Google Scholar
  • View reference in article

• 32

  ChaulagainD.FrugoliJ. (2021). The regulation of nodule number in legumes is a balance of three signal transduction pathways. Int. J. Mol. Sci.22, 1117. doi: 10.3390/ijms22031117

  • CrossRef
  • Google Scholar
  • View reference in article

• 33

  ChenX.YaoQ.GaoX.JiangC.HarberdN. P.FuX. (2016). Shoot-to-Root mobile transcription factor HY5 coordinates plant carbon and nitrogen acquisition. Curr. Biol.26, 640–646. doi: 10.1016/j.cub.2015.12.066

  • CrossRef
  • Google Scholar
  • View reference in article

• 34

  CordobaE.ShishkovaS.VanceC. P.HernándezG. (2003). Antisense inhibition of NADH glutamate synthase impairs carbon/nitrogen assimilation in nodules of alfalfa (Medicago sativa l.). Plant J.33, 1037–1049. doi: 10.1046/j.1365-313X.2003.01686.x

  • CrossRef
  • Google Scholar
  • View reference in article

• 35

  CrookA. D.SchnabelE. L.FrugoliJ. A. (2016). The systemic nodule number regulation kinase SUNN in medicago truncatula interacts with MtCLV2 and MtCRN. Plant J.88, 108–119. doi: 10.1111/tpj.13234

  • CrossRef
  • Google Scholar
  • View reference in article

• 36

  CrozatY.AvelineA.CosteF.GilletJ. P.DomenachA. M. (1994). Yield performance and seed production pattern of field-grown pea and soybean in relation to n nutrition. Eur. J. Agron.3, 135–144. doi: 10.1016/S1161-0301(14)80119-6

  • CrossRef
  • Google Scholar
  • View reference in article

• 37

  DakoraF. D.AtkinsC. A. (1990). Morphological and structural adaptation of nodules of cowpea to functioning under sub- and supra-ambient oxygen pressure. Planta182, 572–582. doi:10.1007

  • Google Scholar
  • View reference in article

• 38

  D’ApuzzoE.ValkovV. T.ParlatiA.OmraneS.BarbulovaA.SainzM. M.et al. (2015). PII overexpression in lotus japonicus affects nodule activity in permissive low-nitrogen conditions and increases nodule numbers in high nitrogen treated plants. MPMI28, 432–442. doi: 10.1094/MPMI-09-14-0285-R

  • CrossRef
  • Google Scholar
  • View reference in article

• 39

  DaubechB.RemigiP.Doin de MouraG.MarchettiM.PouzetC.AuriacM.-C.et al. (2017). Spatio-temporal control of mutualism in legumes helps spread symbiotic nitrogen fixation. Elife6, e28683. doi: 10.7554/eLife.28683

  • CrossRef
  • Google Scholar
  • View reference in article

• 40

  DayD. A.CopelandL. (1991). Carbon metabolism and compartmentation in nitrogen-fixing legume nodules. Plant Physiol. Biochem.29, 185–201.

  • Google Scholar
  • View reference in article

• 41

  DayD. A.MannixM. (1988). Malate oxidation by soybean nodule mitochondria and the possible consequences for nitrogen fixation. Plant Physiol. Biochem.26, 567–573.

  • Google Scholar
  • View reference in article

• 42

  DelayC.ChapmanK.TaleskiM.WangY.TyagiS.XiongY.et al. (2019). CEP3 levels affect starvation-related growth responses of the primary root. J. Exp. Bot.70, 4763–4774. doi: 10.1093/jxb/erz270

  • CrossRef
  • Google Scholar
  • View reference in article

• 43

  DenisonR. F. (1992). Mathematical modeling of oxygen diffusion and respiration in legume root nodules. Plant Physiol.98, 901–907. doi: 10.1104/pp.98.3.901

  • CrossRef
  • Google Scholar
  • View reference in article

• 44

  DenisonR. F.LayzellD. B. (1991). Measurement of legume nodule respiration and O₂ permeability by noninvasive spectrophotometry of leghemoglobin. Plant Physiol.96, 137–143. doi: 10.1104/pp.96.1.137

  • CrossRef
  • Google Scholar
  • View reference in article

• 45

  diCenzoG. C.CheccucciA.BazzicalupoM.MengoniA.VitiC.DziewitL.et al. (2016). Metabolic modelling reveals the specialization of secondary replicons for niche adaptation in sinorhizobium meliloti. Nat. Commun.7, 12219. doi: 10.1038/ncomms12219

  • CrossRef
  • Google Scholar
  • View reference in article

• 46

  DurandJ. L.SheehyJ. E.MinchinF. R. (1987). Nitrogenase activity, photosynthesis and nodule water potential in soyabean plants experiencing water deprivation. J. Exp. Bot.38, 311–321. doi: 10.1093/jxb/38.2.311

  • CrossRef
  • Google Scholar
  • View reference in article

• 47

  DusengeM. E.DuarteA. G.WayD. A. (2019). Plant carbon metabolism and climate change: elevated CO2 and temperature impacts on photosynthesis, photorespiration and respiration. New Phytologist221, 32–49. doi: 10.1111/nph.15283

  • CrossRef
  • Google Scholar
  • View reference in article

• 48

  EvelandA. L.JacksonD. P. (2012). Sugars, signalling, and plant development. J. Exp. Bot.63, 3367–3377. doi: 10.1093/jxb/err379

  • CrossRef
  • Google Scholar
  • View reference in article

• 49

  FedorovaM.TikhonovichI. A.VanceC. P. (1999). Expression of c-assimilating enzymes in pea (Pisum sativum l.) root nodules. In situ localization in effective nodules. Plant Cell Environ.22, 1249–1262. doi: 10.1046/j.1365-3040.1999.00490.x

  • CrossRef
  • Google Scholar
  • View reference in article

• 50

  FergusonB.LinM.-H.GresshoffP. M. (2013). Regulation of legume nodulation by acidic growth conditions. Plant Signaling Behav.8, e23426. doi: 10.4161/psb.23426

  • CrossRef
  • Google Scholar
  • View reference in article

• 51

  FergusonB. J.MensC.HastwellA. H.ZhangM.SuH.JonesC. H.et al. (2019). Legume nodulation: The host controls the party. Plant Cell Environ.42, 41–51. doi: 10.1111/pce.13348

  • CrossRef
  • Google Scholar
  • View reference in article

• 52

  FordeB. G. (2002). Local and long-range signaling pathways regulating plant responses to NO3-. Annu. Rev. Plant Biol.53, 203–224. doi: 10.1146/annurev.arplant.53.100301.135256

  • CrossRef
  • Google Scholar
  • View reference in article

• 53

  FukudomeM.Calvo-BegueriaL.KadoT.OsukiK.RubioM. C.MurakamiE.et al. (2016). Hemoglobin LjGlb1-1 is involved in nodulation and regulates the level of nitric oxide in the lotus japonicus–mesorhizobium loti symbiosis. J. Exp. Bot.67, 5275–5283. doi: 10.1093/jxb/erw290

  • CrossRef
  • Google Scholar
  • View reference in article

• 54

  FukudomeM.WatanabeE.OsukiK.-I.ImaizumiR.AokiT.BecanaM.et al. (2019). Stably transformed lotus japonicus plants overexpressing phytoglobin LjGlb1-1 show decreased nitric oxide levels in roots and nodules as well as delayed nodule senescence. Plant Cell Physiol.60, 816–825. doi: 10.1093/pcp/pcy245

  • CrossRef
  • Google Scholar
  • View reference in article

• 55

  GálvezL.GonzálezE. M.Arrese-IgorC. (2005). Evidence for carbon flux shortage and strong carbon/nitrogen interactions in pea nodules at early stages of water stress. J. Exp. Bot.56, 2551–2561. doi: 10.1093/jxb/eri249

  • CrossRef
  • Google Scholar
  • View reference in article

• 56

  GanY.StulenI.Van KeulenH.KuiperP. J. C. (2002). Physiological changes in soybean (Glycine max) Wuyin9 in response to n and p nutrition. Ann. Appl. Biol.140, 319–329. doi: 10.1111/j.1744-7348.2002.tb00188.x

  • CrossRef
  • Google Scholar
  • View reference in article

• 57

  GangappaS. N.BottoJ. F. (2016). The multifaceted roles of HY5 in plant growth and development. Mol. Plant9, 1353–1365. doi: 10.1016/j.molp.2016.07.002

  • CrossRef
  • Google Scholar
  • View reference in article

• 58

  GanselX.MuñosS.TillardP.GojonA. (2001). Differential regulation of the NO₃^- and NH₄⁺ transporter genes AtNrt2.1 and AtAmt1.1 in arabidopsis: Relation with long-distance and local controls by n status of the plant. Plant J.26, 143–155. doi: 10.1046/j.1365-313x.2001.01016.x

  • CrossRef
  • Google Scholar
  • View reference in article

• 59

  Gaudioso-PedrazaR.BeckM.FrancesL.KirkP.RipodasC.NiebelA.et al. (2018). Callose-regulated symplastic communication coordinates symbiotic root nodule development. Curr. Biol.28, 3562–3577.e6. doi: 10.1016/j.cub.2018.09.031

  • CrossRef
  • Google Scholar
  • View reference in article

• 60

  GautratP.LaffontC.FrugierF. (2020). Compact root architecture 2 promotes root competence for nodulation through the miR2111 systemic effector. Curr. Biol.30, 1339–1345.e3. doi: 10.1016/j.cub.2020.01.084

  • CrossRef
  • Google Scholar
  • View reference in article

• 61

  GautratP.LaffontC.FrugierF.RuffelS. (2021). Nitrogen systemic signaling: From symbiotic nodulation to root acquisition. Trends Plant Sci.26, 392–406. doi: 10.1016/j.tplants.2020.11.009

  • CrossRef
  • Google Scholar
  • View reference in article

• 62

  GautratP.MortierV.LaffontC.De KeyserA.FromentinJ.FrugierF.et al. (2019). Unraveling new molecular players involved in the autoregulation of nodulation in medicago truncatula. J. Exp. Bot.70, 1407–1417. doi: 10.1093/jxb/ery465

  • CrossRef
  • Google Scholar
  • View reference in article

• 63

  Gil-QuintanaE.LarrainzarE.Arrese-IgorC.GonzálezE. M. (2013a). Is n-feedback involved in the inhibition of nitrogen fixation in drought-stressed medicago truncatula? J. Exp. Bot.64, 281–292. doi: 10.1093/jxb/ers334

  • CrossRef
  • Google Scholar
  • View reference in article

• 64

  Gil-QuintanaE.LarrainzarE.SeminarioA.Díaz-LealJ. L.AlamilloJ. M.PinedaM.et al. (2013b). Local inhibition of nitrogen fixation and nodule metabolism in drought-stressed soybean. J. Exp. Bot.64, 2171–2182. doi: 10.1093/jxb/ert074

  • CrossRef
  • Google Scholar
  • View reference in article

• 65

  GirinT.El-KafafiE.-S.WidiezT.ErbanA.HubbertenH.-M.KopkaJ.et al. (2010). Identification of arabidopsis mutants impaired in the systemic regulation of root NO3- uptake by the nitrogen status of the plant. Plant Physiol.153, 1250–1260. doi: 10.1104/pp.110.157354

  • CrossRef
  • Google Scholar
  • View reference in article

• 66

  GirinT.LejayL.WirthJ.WidiezT.PalencharP. M.NazoaP.et al. (2007). Identification of a 150 bp cis-acting element of the AtNRT2.1 promoter involved in the regulation of gene expression by the n and c status of the plant. Plant Cell Environ.30, 1366–1380. doi: 10.1111/j.1365-3040.2007.01712.x

  • CrossRef
  • Google Scholar
  • View reference in article

• 67

  GojonA.CassanO.BachL.LejayL.MartinA. (2022). The decline of plant mineral nutrition under rising CO2: physiological and molecular aspects of a bad deal. Trends Plant Sci.28, 185–198. doi: 10.1016/j.tplants.2022.09.002

  • CrossRef
  • Google Scholar
  • View reference in article

• 68

  GojonA.NacryP.DavidianJ.-C. (2009). Root uptake regulation: A central process for NPS homeostasis in plants. Curr. Opin. Plant Biol.12, 328–338. doi: 10.1016/j.pbi.2009.04.015

  • CrossRef
  • Google Scholar
  • View reference in article

• 69

  GonzálezE. M.Aparicio-TejoP. M.GordonA. J.MinchinF. R.RoyuelaM.Arrese-IgorC. (1998). Water-deficit effects on carbon and nitrogen metabolism of pea nodules. J. Exp. Bot.49, 1705–1714. doi: 10.1093/jxb/49.327.1705

  • CrossRef
  • Google Scholar
  • View reference in article

• 70

  GonzálezE. M.GordonA. J.JamesC. L.Arrese-lgorC. (1995). The role of sucrose synthase in the response of soybean nodules to drought. J. Exp. Bot.46, 1515–1523. doi: 10.1093/jxb/46.10.1515

  • CrossRef
  • Google Scholar
  • View reference in article

• 71

  GordonA. J.MinchinF. R.JamesC. L.KominaO. (1999). Sucrose synthase in legume nodules is essential for nitrogen fixation. Plant Physiol.120, 867–878. doi: 10.1104/pp.120.3.867

  • CrossRef
  • Google Scholar
  • View reference in article

• 72

  GordonA. J.RyleG. J. A.MitchellD. F.PowellD. C. E. (1985). The flux of 14C-labelled photosynthate through soyabean root nodules during N2 fixation. J. Exp. Bot.36, 756–769. doi: 10.1093/jxb/36.5.756

  • CrossRef
  • Google Scholar
  • View reference in article

• 73

  GranotD.David-SchwartzR.KellyG. (2013). Hexose kinases and their role in sugar-sensing and plant development. Front. Plant Sci.4. doi: 10.3389/fpls.2013.00044

  • CrossRef
  • Google Scholar
  • View reference in article

• 74

  GrigstonJ. C.OsunaD.ScheibleW.-R.LiuC.StittM.JonesA. M. (2008). D-glucose sensing by a plasma membrane regulator of G signaling protein, AtRGS1. FEBS Lett.582, 3577–3584. doi: 10.1016/j.febslet.2008.08.038

  • CrossRef
  • Google Scholar
  • View reference in article

• 75

  GrilloM. A.StinchcombeJ. R.HeathK. D. (2016). Nitrogen addition does not influence pre-infection partner choice in the legume-rhizobium symbiosis. Am. J. Bot.103, 1763–1770. doi: 10.3732/ajb.1600090

  • CrossRef
  • Google Scholar
  • View reference in article

• 76

  GuanP.RipollJ.-J.WangR.VuongL.Bailey-SteinitzL. J.YeD.et al. (2017). Interacting TCP and NLP transcription factors control plant responses to NO3- availability. Proc. Natl. Acad. Sci.114, 2419–2424. doi: 10.1073/pnas.1615676114

  • CrossRef
  • Google Scholar
  • View reference in article

• 77

  GuoH.SunY.LiY.LiuX.RenQ.Zhu-SalzmanK.et al. (2013). Elevated CO₂ modifies n acquisition of medicago truncatula by enhancing n fixation and reducing NO₃^- uptake from soil. PloS One8, e81373. doi: 10.1371/journal.pone.0081373

  • CrossRef
  • Google Scholar
  • View reference in article

• 78

  HirschA. M. (1992). Developmental biology of legume nodulation. New Phytol.122, 211–237. doi: 10.1111/j.1469-8137.1992.tb04227.x

  • CrossRef
  • Google Scholar
  • View reference in article

• 79

  HoC.-H.LinS.-H.HuH.-C.TsayY.-F. (2009). CHL1 functions as a NO₃^- sensor in plants. Cell138, 1184–1194. doi: 10.1016/j.cell.2009.07.004

  • CrossRef
  • Google Scholar
  • View reference in article

• 80

  HohnjecN.PerlickA. M.PühlerA.KüsterH. (2003). The medicago truncatula sucrose synthase gene MtSucS1 is activated both in the infected region of root nodules and in the cortex of roots colonized by arbuscular mycorrhizal fungi. MPMI16, 903–915. doi: 10.1094/MPMI.2003.16.10.903

  • CrossRef
  • Google Scholar
  • View reference in article

• 81

  HorchaniF.PrévotM.BoscariA.EvangelistiE.MeilhocE.BruandC.et al. (2011). Both plant and bacterial NO₃^- reductases contribute to nitric oxide production in medicago truncatula nitrogen-fixing nodules. Plant Physiol.155, 1023–1036. doi: 10.1104/pp.110.166140

  • CrossRef
  • Google Scholar
  • View reference in article

• 82

  HuaultE.LaffontC.WenJ.MysoreK. S.RatetP.DucG.et al. (2014). Local and systemic regulation of plant root system architecture and symbiotic nodulation by a receptor-like kinase. PLoS Genet.10, e1004891. doi: 10.1371/journal.pgen.1004891

  • CrossRef
  • Google Scholar
  • View reference in article

• 83

  HuntS.LayzellD. B. (1993). Gas exchange of legume nodules and the regulation of nitrogenase activity. Annu. Rev. Plant Physiol. Plant Mol. Biol.44, 483–511. doi: 10.1146/annurev.pp.44.060193.002411

  • CrossRef
  • Google Scholar
  • View reference in article

• 84

  IgamberdievA. U.BykovaN. V.HillR. D. (2006). Nitric oxide scavenging by barley hemoglobin is facilitated by a monodehydroascorbate reductase-mediated ascorbate reduction of methemoglobin. Planta223, 1033–1040. doi: 10.1007/s00425-005-0146-3

  • CrossRef
  • Google Scholar
  • View reference in article

• 85

  ImsandeJ. (1986). Inhibition of nodule development in soybean by NO₃^- or reduced nitrogen. J. Exp. Bot.37, 348–355. doi: 10.1093/jxb/37.3.348

  • CrossRef
  • Google Scholar
  • View reference in article

• 86

  ImsandeJ.TouraineB. (1994). N demand and the regulation of NO₃^- uptake. Plant Physiol.105, 3–7. doi: 10.1104/pp.105.1.3

  • CrossRef
  • Google Scholar
  • View reference in article

• 87

  JamesE. K.SprentJ. I.MinchinF. R.BrewinN. J. (1991). Intercellular location of glycoprotein in soybean nodules: effect of altered rhizosphere oxygen concentration. Plant Cell Environ.14, 467–476. doi: 10.1111/j.1365-3040.1991.tb01516.x

  • CrossRef
  • Google Scholar
  • View reference in article

• 88

  JangJ. C.LeónP.ZhouL.SheenJ. (1997). Hexokinase as a sugar sensor in higher plants. Plant Cell9, 5–19. doi: 10.1105/tpc.9.1.5

  • CrossRef
  • Google Scholar
  • View reference in article

• 89

  JeudyC.RuffelS.FreixesS.TillardP.SantoniA. L.MorelS.et al. (2010). Adaptation of medicago truncatula to nitrogen limitation is modulated via local and systemic nodule developmental responses. New Phytol.185, 817–828. doi: 10.1111/j.1469-8137.2009.03103.x

  • CrossRef
  • Google Scholar
  • View reference in article

• 90

  JuhnkeH.KremsB.KötterP.EntianK.-D. (1996). Mutants that show increased sensitivity to hydrogen peroxide reveal an important role for the pentose phosphate pathway in protection of yeast against oxidative stress. Molec. Gen. Genet.252, 456–464. doi: 10.1007/BF02173011

  • CrossRef
  • Google Scholar
  • View reference in article

• 91

  KassawT.BridgesW.FrugoliJ. (2015). Multiple autoregulation of nodulation (AON) signals identified through split-root analysis of medicago truncatula sunn and rdn1 mutants. Plants (Basel)4, 209–224. doi: 10.3390/plants4020209

  • CrossRef
  • Google Scholar
  • View reference in article

• 92

  KiersE. T.RousseauR. A.WestS. A.DenisonR. F. (2003). Host sanctions and the legume-rhizobium mutualism. Nature425, 78–81. doi: 10.1038/nature01931

  • CrossRef
  • Google Scholar
  • View reference in article

• 93

  KosslakR. M.BohloolB. B. (1984). Suppression of nodule development of one side of a split-root system of soybeans caused by prior inoculation of the other side. Plant Physiol.75, 125–130. doi: 10.1104/pp.75.1.125

  • CrossRef
  • Google Scholar
  • View reference in article

• 94

  KosslakR. M.BohloolB. B.DowdleS.SadowskyM. J. (1983). Competition of rhizobium japonicum strains in early stages of soybean nodulation. Appl. Environ. Microbiol.46, 870–873. doi: 10.1128/aem.46.4.870-873.1983

  • CrossRef
  • Google Scholar
  • View reference in article

• 95

  KouchiH.FukaiK.KatagiriH.MinamisawaK.TajimaS. (1988). Isolation and enzymological characterization of infected and uninfected cell protoplasts from root nodules of glycine max. Physiologia Plantarum73, 327–334. doi: 10.1111/j.1399-3054.1988.tb00606.x

  • CrossRef
  • Google Scholar
  • View reference in article

• 96

  KroukG.LacombeB.BielachA.Perrine-WalkerF.MalinskaK.MounierE.et al. (2010). NO3–regulated auxin transport by NRT1.1 defines a mechanism for nutrient sensing in plants. Dev. Cell18, 927–937. doi: 10.1016/j.devcel.2010.05.008

  • CrossRef
  • Google Scholar
  • View reference in article

• 97

  KrusellL.MadsenL. H.SatoS.AubertG.GenuaA.SzczyglowskiK.et al. (2002). Shoot control of root development and nodulation is mediated by a receptor-like kinase. Nature420, 422–426. doi: 10.1038/nature01207

  • CrossRef
  • Google Scholar
  • View reference in article

• 98

  KrusellL.SatoN.FukuharaI.KochB. E. V.GrossmannC.OkamotoS.et al. (2011). The Clavata2 genes of pea and lotus japonicus affect autoregulation of nodulation. Plant J.65, 861–871. doi: 10.1111/j.1365-313X.2010.04474.x

  • CrossRef
  • Google Scholar
  • View reference in article

• 99

  KryvoruchkoI. S.SinharoyS.Torres-JerezI.SossoD.PislariuC. I.GuanD.et al. (2016). MtSWEET11, a nodule-specific sucrose transporter of medicago truncatula1[OPEN]. Plant Physiol.171, 554–565. doi: 10.1104/pp.15.01910

  • CrossRef
  • Google Scholar
  • View reference in article

• 100

  LaffontC.HuaultE.GautratP.EndreG.KaloP.BourionV.et al. (2019). Independent regulation of symbiotic nodulation by the SUNN negative and CRA2 positive systemic pathways. Plant Physiol.180, 559–570. doi: 10.1104/pp.18.01588

  • CrossRef
  • Google Scholar
  • View reference in article

• 101

  LaffontC.IvanoviciA.GautratP.BraultM.DjordjevicM. A.FrugierF.et al. (2020). The NIN transcription factor coordinates CEP and CLE signaling peptides that regulate nodulation antagonistically. Nat. Commun.11, 3167. doi: 10.1038/s41467-020-16968-1

  • CrossRef
  • Google Scholar
  • View reference in article

• 102

  LaguerreG.Heulin-GottyK.BrunelB.KlonowskaA.Le QuéréA.TillardP.et al. (2012). Local and systemic n signaling are involved in medicago truncatula preference for the most efficient sinorhizobium symbiotic partners. New Phytol.195, 437–449. doi: 10.1111/j.1469-8137.2012.04159.x

  • CrossRef
  • Google Scholar
  • View reference in article

• 103

  LamS. K.ChenD.NortonR.ArmstrongR.MosierA. R. (2012). Nitrogen dynamics in grain crop and legume pasture systems under elevated atmospheric carbon dioxide concentration: A meta-analysis. Global Change Biol.18, 2853–2859. doi: 10.1111/j.1365-2486.2012.02758.x

  • CrossRef
  • Google Scholar
  • View reference in article

• 104

  LambertI.PerventM.Le QueréA.ClémentG.TauzinM.SeveracD.et al. (2020)a. Responses of mature symbiotic nodules to the whole-plant systemic nitrogen signaling. J. Exp. Bot.71, 5039–5052. doi: 10.1093/jxb/eraa221

  • CrossRef
  • Google Scholar
  • View reference in article

• 105

  LarrainzarE.WienkoopS.ScherlingC.KempaS.LadreraR.Arrese-IgorC.et al. (2009). Carbon metabolism and bacteroid functioning are involved in the regulation of nitrogen fixation in medicago truncatula under drought and recovery. MPMI22, 1565–1576. doi: 10.1094/MPMI-22-12-1565

  • CrossRef
  • Google Scholar
  • View reference in article

• 106

  LebedevaM.AzarakhshM.YashenkovaY.LutovaL. (2020). NO3–induced CLE peptide systemically inhibits nodulation in medicago truncatula. Plants (Basel)9, E1456. doi: 10.3390/plants9111456

  • CrossRef
  • Google Scholar
  • View reference in article

• 107

  LejayL.GanselX.CerezoM.TillardP.MüllerC.KrappA.et al. (2003). Regulation of root ion transporters by photosynthesis: functional importance and relation with hexokinase. Plant Cell15, 2218–2232. doi: 10.1105/tpc.013516

  • CrossRef
  • Google Scholar
  • View reference in article

• 108

  LejayL.WirthJ.PerventM.CrossJ. M.-F.TillardP.GojonA. (2008). Oxidative pentose phosphate pathway-dependent sugar sensing as a mechanism for regulation of root ion transporters by photosynthesis. Plant Physiol.146, 2036–2053. doi: 10.1104/pp.107.114710

  • CrossRef
  • Google Scholar
  • View reference in article

• 109

  LiY.YuZ.LiuX.MathesiusU.WangG.TangC.et al. (2017). Elevated CO2 increases nitrogen fixation at the reproductive phase contributing to various yield responses of soybean cultivars. Front. Plant Sci.8. doi: 10.3389/fpls.2017.01546

  • CrossRef
  • Google Scholar
  • View reference in article

• 110

  LinJ.LiX.LuoZ.MysoreK. S.WenJ.XieF. (2018). NIN interacts with NLPs to mediate NO3- inhibition of nodulation in medicago truncatula. Nat. Plants4, 942–952. doi: 10.1038/s41477-018-0261-3

  • CrossRef
  • Google Scholar
  • View reference in article

• 111

  LindströmK.MousaviS. A. (2020). Effectiveness of nitrogen fixation in rhizobia. Microb. Biotechnol.13, 1314–1335. doi: 10.1111/1751-7915.13517

  • CrossRef
  • Google Scholar
  • View reference in article

• 112

  LiuA.ContadorC. A.FanK.LamH.-M. (2018). Interaction and regulation of carbon, nitrogen, and phosphorus metabolisms in root nodules of legumes. Front. Plant Sci.9. doi: 10.3389/fpls.2018.01860

  • CrossRef
  • Google Scholar
  • View reference in article

• 113

  LiuK.-H.LiuM.LinZ.WangZ.-F.ChenB.LiuC.et al. (2022). NIN-like protein 7 transcription factor is a plant NO3- sensor. Science377, 1419–1425. doi: 10.1126/science.add1104

  • CrossRef
  • Google Scholar
  • View reference in article

• 114

  LiuK.NiuY.KonishiM.WuY.DuH.Sun ChungH.et al. (2017). Discovery of NO₃^–CPK-NLP signalling in central nutrient-growth networks. Nature545, 311–316. doi: 10.1038/nature22077

  • CrossRef
  • Google Scholar
  • View reference in article

• 115

  LiuY.WuL.BaddeleyJ. A.WatsonC. A. (2011). Models of biological nitrogen fixation of legumes. a review. Agron. Sust. Developm.31, 155–172. doi: 10.1051/agro/2010008

  • CrossRef
  • Google Scholar
  • View reference in article

• 116

  LodwigE. M.LeonardM.MarroquiS.WheelerT. R.FindlayK.DownieJ. A.et al. (2005). Role of polyhydroxybutyrate and glycogen as carbon storage compounds in pea and bean bacteroids. MPMI18, 67–74. doi: 10.1094/MPMI-18-0067

  • CrossRef
  • Google Scholar
  • View reference in article

• 117

  LunnJ. E.DelorgeI.FigueroaC. M.Van DijckP.StittM. (2014). Trehalose metabolism in plants. Plant J.79, 544–567. doi: 10.1111/tpj.12509

  • CrossRef
  • Google Scholar
  • View reference in article

• 118

  MaghiaouiA.GojonA.BachL. (2020). NRT1.1-centered NO₃^- signaling in plants. J. Exp. Bot.71, 6226–6237. doi: 10.1093/jxb/eraa361

  • CrossRef
  • Google Scholar
  • View reference in article

• 119

  MagoriS.Oka-KiraE.ShibataS.UmeharaY.KouchiH.HaseY.et al. (2009). Too much love, a root regulator associated with the long-distance control of nodulation in lotus japonicus. Mol. Plant Microbe Interact.22, 259–268. doi: 10.1094/MPMI-22-3-0259

  • CrossRef
  • Google Scholar
  • View reference in article

• 120

  MarchiveC.RoudierF.CastaingsL.BréhautV.BlondetE.ColotV.et al. (2013). Nuclear retention of the transcription factor NLP7 orchestrates the early response to NO3- in plants. Nat. Commun.4, 1713. doi: 10.1038/ncomms2650

  • CrossRef
  • Google Scholar
  • View reference in article

• 121

  MarcoD. E.CarbajalJ. P.CannasS.Pérez-ArnedoR.Hidalgo-PereaA.OlivaresJ.et al. (2009). An experimental and modelling exploration of the host-sanction hypothesis in legume-rhizobia mutualism. J. Theor. Biol.259, 423–433. doi: 10.1016/j.jtbi.2009.03.033

  • CrossRef
  • Google Scholar
  • View reference in article

• 122

  MarinoD.AndrioE.DanchinE. G. J.OgerE.GucciardoS.LambertA.et al. (2011). A medicago truncatula NADPH oxidase is involved in symbiotic nodule functioning. New Phytol.189, 580–592. doi: 10.1111/j.1469-8137.2010.03509.x

  • CrossRef
  • Google Scholar
  • View reference in article

• 123

  MarinoD.DamianiI.GucciardoS.MijangosI.PaulyN.PuppoA. (2013). Inhibition of nitrogen fixation in symbiotic medicago truncatula upon cd exposure is a local process involving leghaemoglobin. J. Exp. Bot.64, 5651–5660. doi: 10.1093/jxb/ert334

  • CrossRef
  • Google Scholar
  • View reference in article

• 124

  MathewsA.CarrollB. J.GresshoffP. M. (1989). Development ofBradyrhizobium infections in supernodulating and non-nodulating mutants of soybean (Glycine max [L.] Merrill). Protoplasma150, 40–47. doi: 10.1007/BF01352919

  • CrossRef
  • Google Scholar
  • View reference in article

• 125

  MensC.HastwellA. H.SuH.GresshoffP. M.MathesiusU.FergusonB. J. (2020). Characterisation of medicago truncatula CLE34 and CLE35 in NO₃^- and rhizobia regulation of nodulation. New Phytol229, 2525–2534. doi: 10.1111/nph.17010

  • CrossRef
  • Google Scholar
  • View reference in article

• 126

  MinchinF. R.WittyJ. F. (2005). “Respiratory/Carbon costs of symbiotic nitrogen fixation in legumes,” in Plant respiration: From cell to ecosystem advances in photosynthesis and respiration. Eds. LambersH.Ribas-CarboM. (Dordrecht: Springer Netherlands), 195–205. doi: 10.1007/1-4020-3589-6_11

  • CrossRef
  • Google Scholar
  • View reference in article

• 127

  MiyazawaH.Oka-KiraE.SatoN.TakahashiH.WuG.-J.SatoS.et al. (2010). The receptor-like kinase KLAVIER mediates systemic regulation of nodulation and non-symbiotic shoot development in lotus japonicus. Development137, 4317–4325. doi: 10.1242/dev.058891

  • CrossRef
  • Google Scholar
  • View reference in article

• 128

  MoreauD.BurstinJ.AubertG.HuguetT.BenC.ProsperiJ.-M.et al. (2012). Using a physiological framework for improving the detection of quantitative trait loci related to nitrogen nutrition in medicago truncatula. Theor. Appl. Genet.124, 755–768. doi: 10.1007/s00122-011-1744-z

  • CrossRef
  • Google Scholar
  • View reference in article

• 129

  MoreauC.GautratP.FrugierF. (2021). NO3–induced CLE35 signaling peptides inhibit nodulation through the SUNN receptor and miR2111 repression. Plant Physiol.185, 1216–1228. doi: 10.1093/plphys/kiaa094

  • CrossRef
  • Google Scholar
  • View reference in article

• 130

  MoreauD.VoisinA.-S.SalonC.Munier-JolainN. (2008). The model symbiotic association between medicago truncatula cv. jemalong and rhizobium meliloti strain 2011 leads to n-stressed plants when symbiotic N2 fixation is the main n source for plant growth. J. Exp. Bot.59, 3509–3522. doi: 10.1093/jxb/ern203

  • CrossRef
  • Google Scholar
  • View reference in article

• 131

  MortierV.Den HerderG.WhitfordR.Van de VeldeW.RombautsS.D’HaeseleerK.et al. (2010). CLE peptides control medicago truncatula nodulation locally and systemically. Plant Physiol.153, 222–237. doi: 10.1104/pp.110.153718

  • CrossRef
  • Google Scholar
  • View reference in article

• 132

  MullerB.TouraineB. (1992). Inhibition of NO⊟3 uptake by various phloem-translocated amino acids in soybean seedlings. J. Exp. Bot.43, 617–623. doi: 10.1093/jxb/43.5.617

  • CrossRef
  • Google Scholar
  • View reference in article

• 133

  NayaL.LadreraR.RamosJ.GonzálezE. M.Arrese-IgorC.MinchinF. R.et al. (2007). The response of carbon metabolism and antioxidant defenses of alfalfa nodules to drought stress and to the subsequent recovery of plants. Plant Physiol.144, 1104–1114. doi: 10.1104/pp.107.099648

  • CrossRef
  • Google Scholar
  • View reference in article

• 134

  NeoH. H.LayzellD. B. (1997). Phloem glutamine and the regulation of O₂ diffusion in legume nodules. Plant Physiol.113, 259–267. doi: 10.1104/pp.113.1.259

  • CrossRef
  • Google Scholar
  • View reference in article

• 135

  NeuhausH. E.EmesM. J. (2000). Nonphotosynthetic metabolism in plastids. Annu. Rev. Plant Physiol. Plant Mol. Biol.51, 111–140. doi: 10.1146/annurev.arplant.51.1.111

  • CrossRef
  • Google Scholar
  • View reference in article

• 136

  NishidaH.HandaY.TanakaS.SuzakiT.KawaguchiM. (2016). Expression of the CLE-RS3 gene suppresses root nodulation in lotus japonicus. J. Plant Res.129, 909–919. doi: 10.1007/s10265-016-0842-z

  • CrossRef
  • Google Scholar
  • View reference in article

• 137

  NishidaH.ItoM.MiuraK.KawaguchiM.SuzakiT. (2020). Autoregulation of nodulation pathway is dispensable for NO₃^–induced control of rhizobial infection. Plant Signal Behav.15, 1733814. doi: 10.1080/15592324.2020.1733814

  • CrossRef
  • Google Scholar
  • View reference in article

• 138

  NishidaH.TanakaS.HandaY.ItoM.SakamotoY.MatsunagaS.et al. (2018). A NIN-LIKE PROTEIN mediates NO₃^–induced control of root nodule symbiosis in lotus japonicus. Nat. Commun.9, 499. doi: 10.1038/s41467-018-02831-x

  • CrossRef
  • Google Scholar
  • View reference in article

• 139

  NishimuraR.HayashiM.WuG.-J.KouchiH.Imaizumi-AnrakuH.MurakamiY.et al. (2002). HAR1 mediates systemic regulation of symbiotic organ development. Nature420, 426–429. doi: 10.1038/nature01231

  • CrossRef
  • Google Scholar
  • View reference in article

• 140

  NoctorG.FoyerC. H. (1998). ASCORBATE AND GLUTATHIONE: Keeping active oxygen under control. Annu. Rev. Plant Physiol. Plant Mol. Biol.49, 249–279. doi: 10.1146/annurev.arplant.49.1.249

  • CrossRef
  • Google Scholar
  • View reference in article

• 141

  OhkuboY.TanakaM.TabataR.Ogawa-OhnishiM.MatsubayashiY. (2017). Shoot-to-root mobile polypeptides involved in systemic regulation of nitrogen acquisition. Nat. Plants3, 17029. doi: 10.1038/nplants.2017.29

  • CrossRef
  • Google Scholar
  • View reference in article

• 142

  OjiY.WatanabeM.WakiuchiN.OkamotoS. (1985). Nitrite reduction in barley-root plastids: Dependence on NADPH coupled with glucose-6-phosphate and 6-phosphogluconate dehydrogenases, and possible involvement of an electron carrier and a diaphorase. Planta165, 85–90. doi: 10.1007/BF00392215

  • CrossRef
  • Google Scholar
  • View reference in article

• 143

  OkamotoS.OhnishiE.SatoS.TakahashiH.NakazonoM.TabataS.et al. (2009). Nod factor/NO3–induced CLE genes that drive HAR1-mediated systemic regulation of nodulation. Plant Cell Physiol.50, 67–77. doi: 10.1093/pcp/pcn194

  • CrossRef
  • Google Scholar
  • View reference in article

• 144

  OldroydG. E. D.DownieJ. A. (2008). Coordinating nodule morphogenesis with rhizobial infection in legumes. Annu. Rev. Plant Biol.59, 519–546. doi: 10.1146/annurev.arplant.59.032607.092839

  • CrossRef
  • Google Scholar
  • View reference in article

• 145

  OldroydG. E. D.MurrayJ. D.PooleP. S.DownieJ. A. (2011). The rules of engagement in the legume-rhizobial symbiosis. Annu. Rev. Genet.45, 119–144. doi: 10.1146/annurev-genet-110410-132549

  • CrossRef
  • Google Scholar
  • View reference in article

• 146

  OlssonJ. E.NakaoP.BohloolB. B.GresshoffP. M. (1989). Lack of systemic suppression of nodulation in split-root systems of supernodulating soybean (Glycine max [L.] merr.) mutants. Plant Physiol.90, 1347–1352. doi: 10.1104/pp.90.4.1347

  • CrossRef
  • Google Scholar
  • View reference in article

• 147

  OonoR.AndersonC. G.DenisonR. F. (2011). Failure to fix nitrogen by non-reproductive symbiotic rhizobia triggers host sanctions that reduce fitness of their reproductive clonemates. Proc. Biol. Sci.278, 2698–2703. doi: 10.1098/rspb.2010.2193

  • CrossRef
  • Google Scholar
  • View reference in article

• 148

  OonoR.DenisonR. F. (2010). Comparing symbiotic efficiency between swollen versus nonswollen rhizobial bacteroids. Plant Physiol.154, 1541–1548. doi: 10.1104/pp.110.163436

  • CrossRef
  • Google Scholar
  • View reference in article

• 149

  OonoR.DenisonR. F.KiersE. T. (2009). Controlling the reproductive fate of rhizobia: how universal are legume sanctions? New Phytol.183, 967–979. doi: 10.1111/j.1469-8137.2009.02941.x

  • CrossRef
  • Google Scholar
  • View reference in article

• 150

  ParsonsR.StanforthA.RavenJ. A.SprentJ. I. (1993). Nodule growth and activity may be regulated by a feedback mechanism involving phloem nitrogen. Plant Cell Environ.16, 125–136. doi: 10.1111/j.1365-3040.1993.tb00854.x

  • CrossRef
  • Google Scholar
  • View reference in article

• 151

  ParvinS.UddinS.Tausz-PoschS.ArmstrongR.TauszM. (2020). Carbon sink strength of nodules but not other organs modulates photosynthesis of faba bean (Vicia faba) grown under elevated [CO2] and different water supply. New Phytol.227, 132–145. doi: 10.1111/nph.16520

  • CrossRef
  • Google Scholar
  • View reference in article

• 152

  PatriarcaE. J.TatèR.IaccarinoM. (2002). Key role of bacterial NH₄⁺ metabolism in rhizobium-plant symbiosis. Microbiol. Mol. Biol. Rev.66, 203–222. doi: 10.1128/MMBR.66.2.203-222.2002

  • CrossRef
  • Google Scholar
  • View reference in article

• 153

  PaulyN.PucciarielloC.MandonK.InnocentiG.JametA.BaudouinE.et al. (2006). Reactive oxygen and nitrogen species and glutathione: Key players in the legume-rhizobium symbiosis. J. Exp. Bot.57, 1769–1776. doi: 10.1093/jxb/erj184

  • CrossRef
  • Google Scholar
  • View reference in article

• 154

  PenmetsaR. V.FrugoliJ. A.SmithL. S.LongS. R.CookD. R. (2003). Dual genetic pathways controlling nodule number in medicago truncatula. Plant Physiol.131, 998–1008. doi: 10.1104/pp.015677

  • CrossRef
  • Google Scholar
  • View reference in article

• 155

  Pérez GuerraJ. C.CoussensG.De KeyserA.De RyckeR.De BodtS.Van De VeldeW.et al. (2010). Comparison of developmental and stress-induced nodule senescence in medicago truncatula. Plant Physiol.152, 1574–1584. doi: 10.1104/pp.109.151399

  • CrossRef
  • Google Scholar
  • View reference in article

• 156

  PerventM.LambertI.TauzinM.KarouaniA.NiggM.JardinaudM.-F.et al. (2021). Systemic control of nodule formation by plant nitrogen demand requires autoregulation-dependent and independent mechanisms. J. Exp. Bot.72, 7942–7956. doi: 10.1093/jxb/erab374

  • CrossRef
  • Google Scholar
  • View reference in article

• 157

  PessiG.AhrensC. H.RehrauerH.LindemannA.HauserF.FischerH.-M.et al. (2007). Genome-wide transcript analysis of bradyrhizobium japonicum bacteroids in soybean root nodules. MPMI20, 1353–1363. doi: 10.1094/MPMI-20-11-1353

  • CrossRef
  • Google Scholar
  • View reference in article

• 158

  PrayitnoJ.RolfeB. G.MathesiusU. (2006). The ethylene-insensitive sickle mutant of medicago truncatula shows altered auxin transport regulation during nodulation. Plant Physiol.142, 168–180. doi: 10.1104/pp.106.080093

  • CrossRef
  • Google Scholar
  • View reference in article

• 159

  PrellJ.WhiteJ. P.BourdesA.BunnewellS.BongaertsR. J.PooleP. S. (2009). Legumes regulate rhizobium bacteroid development and persistence by the supply of branched-chain amino acids. Proc. Natl. Acad. Sci. U.S.A.106, 12477–12482. doi: 10.1073/pnas.0903653106

  • CrossRef
  • Google Scholar
  • View reference in article

• 160

  PuppoA.PaulyN.BoscariA.MandonK.BrouquisseR. (2013). Hydrogen peroxide and nitric oxide: key regulators of the legume-rhizobium and mycorrhizal symbioses. Antioxid Redox Signal18, 2202–2219. doi: 10.1089/ars.2012.5136

  • CrossRef
  • Google Scholar
  • View reference in article

• 161

  RalserM.WamelinkM. M.KowaldA.GerischB.HeerenG.StruysE. A.et al. (2007). Dynamic rerouting of the carbohydrate flux is key to counteracting oxidative stress. J. Biol.6, 10. doi: 10.1186/jbiol61

  • CrossRef
  • Google Scholar
  • View reference in article

• 162

  ReidD. E.FergusonB. J.GresshoffP. M. (2011a). Inoculation- and NO3–induced CLE peptides of soybean control NARK-dependent nodule formation. Mol. Plant Microbe Interact.24, 606–618. doi: 10.1094/MPMI-09-10-0207

  • CrossRef
  • Google Scholar
  • View reference in article

• 163

  ReidD. E.FergusonB. J.HayashiS.LinY.-H.GresshoffP. M. (2011b). Molecular mechanisms controlling legume autoregulation of nodulation. Ann. Bot.108, 789–795. doi: 10.1093/aob/mcr205

  • CrossRef
  • Google Scholar
  • View reference in article

• 164

  RiverasE.AlvarezJ. M.VidalE. A.OsesC.VegaA.GutiérrezR. A. (2015). The calcium ion is a second messenger in the NO₃^- signaling pathway of arabidopsis. Plant Physiol.169, 1397–1404. doi: 10.1104/pp.15.00961

  • CrossRef
  • Google Scholar
  • View reference in article

• 165

  RobertsonJ. G.WellsB.BisselingT.FarndenK. J. F.JohnstonA. W. B. (1984). Immuno-gold localization of leghaemoglobin in cytoplasm in nitrogen-fixing root nodules of pea. Nature311, 254–256. doi: 10.1038/311254a0

  • CrossRef
  • Google Scholar
  • View reference in article

• 166

  RogersA.GibonY.StittM.MorganP. B.BernacchiC. J.OrtD. R.et al. (2006). Increased c availability at elevated carbon dioxide concentration improves n assimilation in a legume. Plant Cell Environ.29, 1651–1658. doi: 10.1111/j.1365-3040.2006.01549.x

  • CrossRef
  • Google Scholar
  • View reference in article

• 167

  RogersA.AinsworthE. A.LeakeyA. D. B. (2009). Will elevated carbon dioxide concentration amplify the benefits of nitrogen fixation in legumes? Plant Physiol.151, 1009–1016. doi: 10.1104/pp.109.144113

  • CrossRef
  • Google Scholar
  • View reference in article

• 168

  RollandF.Baena-GonzalezE.SheenJ. (2006). Sugar sensing and signaling in plants: conserved and novel mechanisms. Annu. Rev. Plant Biol.57, 675–709. doi: 10.1146/annurev.arplant.57.032905.105441

  • CrossRef
  • Google Scholar
  • View reference in article

• 169

  RonsonC. W.AstwoodP. M.DownieJ. A. (1984). Molecular cloning and genetic organization of C4-dicarboxylate transport genes from rhizobium leguminosarum. J. Bacteriol160, 903–909. doi: 10.1128/jb.160.3.903-909.1984

  • CrossRef
  • Google Scholar
  • View reference in article

• 170

  RosendahlL.VanceC. P.PedersenW. B. (1990). Products of dark CO₂ fixation in pea root nodules support bacteroid metabolism 1. Plant Physiol.93, 12–19. doi: 10.1104/pp.93.1.12

  • CrossRef
  • Google Scholar
  • View reference in article

• 171

  RoyS.LiuW.NandetyR. S.CrookA.MysoreK. S.PislariuC. I.et al. (2020). Celebrating 20 years of genetic discoveries in legume nodulation and symbiotic nitrogen fixation. Plant Cell32, 15–41. doi: 10.1105/tpc.19.00279

  • CrossRef
  • Google Scholar
  • View reference in article

• 172

  RoyS.MüllerL. M. (2022). A rulebook for peptide control of legume-microbe endosymbioses. Trends Plant Sci.27, 870–889. doi: 10.1016/j.tplants.2022.02.002

  • CrossRef
  • Google Scholar
  • View reference in article

• 173

  RuffelS.FreixesS.BalzergueS.TillardP.JeudyC.Martin-MagnietteM. L.et al. (2008). Systemic signaling of the plant nitrogen status triggers specific transcriptome responses depending on the nitrogen source in medicago truncatula. Plant Physiol.146, 2020–2035. doi: 10.1104/pp.107.115667

  • CrossRef
  • Google Scholar
  • View reference in article

• 174

  RuffelS.GojonA.LejayL. (2014). Signal interactions in the regulation of root NO₃^- uptake. J. Exp. Bot.65, 5509–5517. doi: 10.1093/jxb/eru321

  • CrossRef
  • Google Scholar
  • View reference in article

• 175

  SaganM.MorandiD.TarenghiE.DucG. (1995). Selection of nodulation and mycorrhizal mutants in the model plant medicago truncatula (Gaertn.) after γ-ray mutagenesis. Plant Sci.111, 63–71. doi: 10.1016/0168-9452(95)04229-N

  • CrossRef
  • Google Scholar
  • View reference in article

• 176

  SalonC.LepetitM.GamasP.JeudyC.MoreauS.MoreauD.et al. (2009). Analysis and modeling of the integrative response of medicago truncatula to nitrogen constraints. C. R. Biol.332, 1022–1033. doi: 10.1016/j.crvi.2009.09.009

  • CrossRef
  • Google Scholar
  • View reference in article

• 177

  SánchezC.GatesA. J.MeakinG. E.UchiumiT.GirardL.RichardsonD. J.et al. (2010). Production of nitric oxide and nitrosylleghemoglobin complexes in soybean nodules in response to flooding. MPMI23, 702–711. doi: 10.1094/MPMI-23-5-0702

  • CrossRef
  • Google Scholar
  • View reference in article

• 178

  Sanz-SáezÁ.EriceG.AranjueloI.NoguésS.IrigoyenJ. J.Sánchez-DíazM. (2010). Photosynthetic down-regulation under elevated CO₂ exposure can be prevented by nitrogen supply in nodulated alfalfa. J. Plant Physiol.167, 1558–1565. doi: 10.1016/j.jplph.2010.06.015

  • CrossRef
  • Google Scholar
  • View reference in article

• 179

  SasakiT.SuzakiT.SoyanoT.KojimaM.SakakibaraH.KawaguchiM. (2014). Shoot-derived cytokinins systemically regulate root nodulation. Nat. Commun.5, 4983. doi: 10.1038/ncomms5983

  • CrossRef
  • Google Scholar
  • View reference in article

• 180

  SchnabelE.JournetE.-P.de Carvalho-NiebelF.DucG.FrugoliJ. (2005). The medicago truncatula SUNN gene encodes a CLV1-likeLeucine-rich repeat receptor kinase that regulates nodule number and root length. Plant Mol. Biol.58, 809–822. doi: 10.1007/s11103-005-8102-y

  • CrossRef
  • Google Scholar
  • View reference in article

• 181

  SchuizeJ.AdgoE.MerbachW. (1999). Carbon costs associated with N2 fixation in vicia faba l and pisum sativum 1. over a 14-day period. Plant Biol.1, 625–631. doi: 10.1111/j.1438-8677.1999.tb00273.x

  • CrossRef
  • Google Scholar
  • View reference in article

• 182

  SchulteC. C. M.BorahK.WheatleyR. M.TerpolilliJ. J.SaalbachG.CrangN.et al. (2021). Metabolic control of nitrogen fixation in rhizobium-legume symbioses. Sci. Adv.7, eabh2433. doi: 10.1126/sciadv.abh2433

  • CrossRef
  • Google Scholar
  • View reference in article

• 183

  SchulzeJ. (2004). How are nitrogen fixation rates regulated in legumes? J. Plant Nutr. Soil Sci.167, 125–137. doi: 10.1002/jpln.200320358

  • CrossRef
  • Google Scholar
  • View reference in article

• 184

  SchwemberA. R.SchulzeJ.del PozoA.CabezaR. A. (2019). Regulation of symbiotic nitrogen fixation in legume root nodules. Plants8, 333. doi: 10.3390/plants8090333

  • CrossRef
  • Google Scholar
  • View reference in article

• 185

  SeabraA. R.PereiraP. A.BeckerJ. D.CarvalhoH. G. (2012). Inhibition of glutamine synthetase by phosphinothricin leads to transcriptome reprograming in root nodules of medicago truncatula. Mol. Plant Microbe Interact.25, 976–992. doi: 10.1094/MPMI-12-11-0322

  • CrossRef
  • Google Scholar
  • View reference in article

• 186

  SearleI. R.MenA. E.LaniyaT. S.BuzasD. M.Iturbe-OrmaetxeI.CarrollB. J.et al. (2003). Long-distance signaling in nodulation directed by a CLAVATA1-like receptor kinase. Science299, 109–112. doi: 10.1126/science.1077937

  • CrossRef
  • Google Scholar
  • View reference in article

• 187

  SingletonP. W.van KesselC. (1987). Effect of localized nitrogen availability to soybean half-root systems on photosynthate partitioning to roots and nodules. Plant Physiol.83, 552–556. doi: 10.1104/pp.83.3.552

  • CrossRef
  • Google Scholar
  • View reference in article

• 188

  SmithP. M. C.WinterH.StorerP. J.BussellJ. D.SchullerK. A.AtkinsC. A. (2002). Effect of short-term N₂ deficiency on expression of the ureide pathway in cowpea root nodules. Plant Physiol.129, 1216–1221. doi: 10.1104/pp.010714

  • CrossRef
  • Google Scholar
  • View reference in article

• 189

  SoupèneE.FoussardM.BoistardP.TruchetG.BatutJ. (1995). Oxygen as a key developmental regulator of rhizobium meliloti N₂-fixation gene expression within the alfalfa root nodule. Proc. Natl. Acad. Sci. U.S.A.92, 3759–3763. doi: 10.1073/pnas.92.9.3759

  • CrossRef
  • Google Scholar
  • View reference in article

• 190

  SprentJ. I. (2009). “Development and functioning of nodules,” in Legume nodulation (John Wiley & Sons, Ltd), 79–95. doi: 10.1002/9781444316384.ch5

  • CrossRef
  • Google Scholar
  • View reference in article

• 191

  StittM.KrappA. (1999). The interaction between elevated carbon dioxide and nitrogen nutrition: the physiological and molecular background. Plant Cell Environ.22, 583–621. doi: 10.1046/j.1365-3040.1999.00386.x

  • CrossRef
  • Google Scholar
  • View reference in article

• 192

  StreeterJ.WongP. P. (1988). Inhibition of legume nodule formation and N2 fixation by NO3-. Crit. Rev. Plant Sci.7, 1–23. doi: 10.1080/07352688809382257

  • CrossRef
  • Google Scholar
  • View reference in article

• 193

  SugiyamaA.SaidaY.YoshimizuM.TakanashiK.SossoD.FrommerW. B.et al. (2017). Molecular characterization of LjSWEET3, a sugar transporter in nodules of lotus japonicus. Plant Cell Physiol.58, 298–306. doi: 10.1093/pcp/pcw190

  • CrossRef
  • Google Scholar
  • View reference in article

• 194

  TabataR.SumidaK.YoshiiT.OhyamaK.ShinoharaH.MatsubayashiY. (2014). Perception of root-derived peptides by shoot LRR-RKs mediates systemic n-demand signaling. Science346, 343–346. doi: 10.1126/science.1257800

  • CrossRef
  • Google Scholar
  • View reference in article

• 195

  TakaharaM.MagoriS.SoyanoT.OkamotoS.YoshidaC.YanoK.et al. (2013). Too much love, a novel kelch repeat-containing f-box protein, functions in the long-distance regulation of the legume-rhizobium symbiosis. Plant Cell Physiol.54, 433–447. doi: 10.1093/pcp/pct022

  • CrossRef
  • Google Scholar
  • View reference in article

• 196

  TakanashiK.SasakiT.KanT.SaidaY.SugiyamaA.YamamotoY.et al. (2016). A dicarboxylate transporter, LjALMT4, mainly expressed in nodules of lotus japonicus. MPMI29, 584–592. doi: 10.1094/MPMI-04-16-0071-R

  • CrossRef
  • Google Scholar
  • View reference in article

• 197

  TakanashiK.TakahashiH.SakuraiN.SugiyamaA.SuzukiH.ShibataD.et al. (2012). Tissue-specific transcriptome analysis in nodules of lotus japonicus. MPMI25, 869–876. doi: 10.1094/MPMI-01-12-0011-R

  • CrossRef
  • Google Scholar
  • View reference in article

• 198

  TerpolilliJ. J.MasakapalliS. K.KarunakaranR.WebbI. U. C.GreenR.WatmoughN. J.et al. (2016). Lipogenesis and redox balance in nitrogen-fixing pea bacteroids. J. Bacteriol198, 2864–2875. doi: 10.1128/JB.00451-16

  • CrossRef
  • Google Scholar
  • View reference in article

• 199

  ThumfortP. P.AtkinsC. A.LayzellD. B. (1994). A re-evaluation of the role of the infected cell in the control of O₂ diffusion inlegume nodules. Plant Physiol.105, 1321–1333. doi: 10.1104/pp.105.4.1321

  • CrossRef
  • Google Scholar
  • View reference in article

• 200

  ThumfortP. P.LayzellD. B.AtkinsC. A. (1999). Diffusion and reaction of oxygen in the central tissue ofureide-producing legume nodules. Plant Cell Environ.22, 1351–1363. doi: 10.1046/j.1365-3040.1999.00498.x

  • CrossRef
  • Google Scholar
  • View reference in article

• 201

  ThumfortP. P.LayzellD. B.AtkinsC. A. (2000). A simplified approach for modeling diffusion into cells. J. Theor. Biol.204, 47–65. doi: 10.1006/jtbi.2000.1071

  • CrossRef
  • Google Scholar
  • View reference in article

• 202

  TillardP.PassamaL.GojonA. (1998). Are phloem amino acids involved in the shoot to root control of NO-3 uptake in ricinus communis plants? J. Exp. Bot.49, 1371–1379. doi: 10.1093/jxb/49.325.1371

  • CrossRef
  • Google Scholar
  • View reference in article

• 203

  TimmersA. C.SoupèneE.AuriacM. C.de BillyF.VasseJ.BoistardP.et al. (2000). Saprophytic intracellular rhizobia in alfalfa nodules. Mol. Plant Microbe Interact.13, 1204–1213. doi: 10.1094/MPMI.2000.13.11.1204

  • CrossRef
  • Google Scholar
  • View reference in article

• 204

  TjepkemaJ. D.YocumC. S. (1974). Measurement of oxygen partial pressure within soybean nodules by oxygen microelectrodes. Planta119, 351–360. doi: 10.1007/BF00388335

  • CrossRef
  • Google Scholar
  • View reference in article

• 205

  TsikouD.YanZ.HoltD. B.AbelN. B.ReidD. E.MadsenL. H.et al. (2018). Systemic control of legume susceptibility to rhizobial infection by a mobile microRNA. Science362, 233–236. doi: 10.1126/science.aat6907

  • CrossRef
  • Google Scholar
  • View reference in article

• 206

  UdvardiM.PooleP. S. (2013). Transport and metabolism in legume-rhizobia symbioses. Annu. Rev. Plant Biol.64, 781–805. doi: 10.1146/annurev-arplant-050312-120235

  • CrossRef
  • Google Scholar

• 207

  UranoD.PhanN.JonesJ. C.YangJ.HuangJ.GrigstonJ.et al. (2012). Endocytosis of the seven-transmembrane RGS1 protein activates G-protein-coupled signalling in arabidopsis. Nat. Cell Biol.14, 1079–1088. doi: 10.1038/ncb2568

  • CrossRef
  • Google Scholar

• 208

  ValkovV. T.RogatoA.AlvesL. M.SolS.NogueroM.LéranS.et al. (2017). The nitrate transporter family protein LjNPF8.6 controls the n-fixing nodule activity. Plant Physiol175, 1269–1282. doi: 10.1104/pp.17.01187

  • CrossRef
  • Google Scholar

• 209

  ValkovV. T.SolS.RogatoA.ChiurazziM. (2020). The functional characterization of LjNRT2.4 indicates a novel, positive role of nitrate for an efficient nodule N2 -fixation activity. New Phytol.228, 682–696. doi: 10.1111/nph.16728

  • CrossRef
  • Google Scholar

• 210

  VanceC. P.HeichelG. H. (1991). Carbon in N2 fixation: Limitation or exquisite adaptation. Annu. Rev. Plant Physiol. Plant Mol. Biol.42, 373–390. doi: 10.1146/annurev.pp.42.060191.002105

  • CrossRef
  • Google Scholar

• 211

  VanceC. P. (2008). Carbon and Nitrogen Metabolism in Legume Nodules. in Nitrogen-fixing Leguminous Symbioses Nitrogen Fixation: Origins, Applications, and Research Progress. DilworthM. J.JamesE. K.SprentJ. I.NewtonW. E. (Dordrecht: Springer Netherlands), 293–320. doi: 10.1007/978-1-4020-3548-7_10

  • CrossRef
  • Google Scholar

• 212

  van NoordenG. E.RossJ. J.ReidJ. B.RolfeB. G.MathesiusU. (2006). Defective long-distance auxin transport regulation in the medicago truncatula super numeric nodules mutant. Plant Physiol.140, 1494–1506. doi: 10.1104/pp.105.075879

  • CrossRef
  • Google Scholar

• 213

  VaughnM. W.HarringtonG. N.BushD. R. (2002). Sucrose-mediated transcriptional regulation of sucrose symporter activity in the phloem. Proc. Natl. Acad. Sci.99, 10876–10880. doi: 10.1073/pnas.172198599

  • CrossRef
  • Google Scholar

• 214

  VerhoevenJ. T. A.KoerselmanW.MeulemanA. F. M. (1996). Nitrogen- or phosphorus-limited growth in herbaceous, wet vegetation: Relations with atmospheric inputs and management regimes. Trends Ecol. Evol.11, 494–497. doi: 10.1016/S0169-5347(96)10055-0

  • CrossRef
  • Google Scholar

• 215

  VidalE. A.AlvarezJ. M.ArausV.RiverasE.BrooksM. D.KroukG.et al. (2020). Nitrate in 2020: Thirty years from transport to signaling networks. Plant Cell32, 2094–2119. doi: 10.1105/tpc.19.00748

  • CrossRef
  • Google Scholar

• 216

  VoisinA. S.SalonC.JeudyC.WarembourgF. R. (2003). Symbiotic N₂ fixation activity in relation to c economy of pisum sativum l. as a function of plant phenology. J. Exp. Bot.54, 2733–2744. doi: 10.1093/jxb/erg290

  • CrossRef
  • Google Scholar

• 217

  WalshK. B.VesseyJ. K.LayzellD. B. (1987). Carbohydrate supply and N2 fixation in soybean 1. Plant Physiol.85, 137–144. doi: 10.1104/pp.85.1.137

  • CrossRef
  • Google Scholar

• 218

  WangT.GuoJ.PengY.LyuX.LiuB.SunS.et al. (2021). Light-induced mobile factors from shoots regulate rhizobium-triggered soybean root nodulation. Science374, 65–71. doi: 10.1126/science.abh2890

  • CrossRef
  • Google Scholar

• 219

  WangR.TischnerR.GutiérrezR. A.HoffmanM.XingX.ChenM.et al. (2004). Genomic analysis of the nitrate response using a nitrate reductase-null mutant of arabidopsis. Plant Physiol.136, 2512–2522. doi: 10.1104/pp.104.044610

  • CrossRef
  • Google Scholar

• 220

  WidiezT.El KafafiE. S.GirinT.BerrA.RuffelS.KroukG.et al. (2011). High nitrogen insensitive 9 (HNI9)-mediated systemic repression of root NO3- uptake is associated with changes in histone methylation. Proc. Natl. Acad. Sci. U.S.A.108, 13329–13334. doi: 10.1073/pnas.1017863108

  • CrossRef
  • Google Scholar

• 221

  WopereisJ.PajueloE.DazzoF. B.JiangQ.GresshoffP. M.De BruijnF. J.et al. (2000). Short root mutant of lotus japonicus with a dramatically altered symbiotic phenotype. Plant J.23, 97–114. doi: 10.1046/j.1365-313x.2000.00799.x

  • CrossRef
  • Google Scholar

• 222

  XiaoW.SheenJ.JangJ. C.. (2000). The role of hexokinase in plant sugar signal transduction and growth and development. Plant Mol. Biol.44, 451–61. doi: 10.1023/a:1026501430422

  • CrossRef
  • Google Scholar

• 223

  YamaguchiY. L.IshidaT.SawaS. (2016). CLE peptides and their signaling pathways in plant development. J. Exp. Bot.67, 4813–4826. doi: 10.1093/jxb/erw208

  • CrossRef
  • Google Scholar

• 224

  YamashitaN.TanabataS.OhtakeN.SueyoshiK.SatoT.HiguchiK.et al. (2019). Effects of different chemical forms of nitrogen on the quick and reversible inhibition of soybean nodule growth and nitrogen fixation activity. Front. Plant Sci.10. doi: 10.3389/fpls.2019.00131

  • CrossRef
  • Google Scholar

• 225

  YurgelS. N.KahnM. L. (2004). Dicarboxylate transport by rhizobia. FEMS Microbiol. Rev.28, 489–501. doi: 10.1016/j.femsre.2004.04.002

  • CrossRef
  • Google Scholar

• 226

  YurgelS. N.RiceJ.KahnM. L. (2012). Nitrogen metabolism in sinorhizobium meliloti–alfalfa symbiosis: Dissecting the role of GlnD and PII proteins. MPMI25, 355–362. doi: 10.1094/MPMI-09-11-0249

  • CrossRef
  • Google Scholar

• 227

  ZhangH.JenningsA.BarlowP. W.FordeB. G. (1999). Dual pathways for regulation of root branching by nitrate. Proc. Natl. Acad. Sci.96, 6529–6534. doi: 10.1073/pnas.96.11.6529

  • CrossRef
  • Google Scholar

• 228

  ZhuF.DengJ.ChenH.LiuP.ZhengL.YeQ.et al. (2020). A CEP peptide receptor-like kinase regulates auxin biosynthesis and ethylene signaling to coordinate root growth and symbiotic nodulation in medicago truncatula. Plant Cell32, 2855–2877. doi: 10.1105/tpc.20.00248

  • CrossRef
  • Google Scholar