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. 2025 Jun 13;177(3):e70321. doi: 10.1111/ppl.70321

The Urea Cycle in Connection to Polyamine Metabolism in Higher Plants: New Perspectives on a Central Pathway

J Buezo 1, M Urra 1, E M González 2, R Alcázar 3, D Marino 4, J F Moran 1,✉
PMCID: PMC12163876  PMID: 40511565

ABSTRACT

The ornithine‐urea cycle is a biochemical pathway primarily found in animals, where it plays a crucial role in the re‐assimilation of ammonium and the removal of excess nitrogen in the form of urea. In lower photosynthetic eukaryotes, it contributes to metabolic responses during episodes of high nitrogen availability. In higher plants, although historically overlooked, compelling evidence indicates the pivotal role of the urea cycle in different aspects of plant physiology and metabolism. In particular, it is associated with the metabolism of polyamines during stress. Unlike in animals and lower photosynthetic eukaryotes, in higher plants, the urea cycle is not complete due to the lack of the carbamoyl phosphate synthase‐I enzyme that incorporates ammonium into the cycle. Higher plants only possess a type‐II carbamoyl phosphate synthase‐II that introduces glutamine into the cycle, which is also metabolically linked to arginine and polyamine metabolism. Putrescine accumulation is a metabolic hallmark of different types of abiotic stresses, such as drought, salinity, ammonium stress, iron and phosphorus deficiency, and low temperatures. Notably, the exogenous application of polyamines, such as putrescine or spermine, enhances tolerance to abiotic stress, a process in which the free radical nitric oxide appears to play a role. Overall, this review article attempts to bring together the current knowledge on the functionality of the constituent enzymes and metabolites of the urea cycle and discuss the importance of this pathway in relation to the metabolism of polyamine in higher plants.

Keywords: abiotic stress, ammonium stress, carbamoyl phosphate synthase, nitric oxide, ornithine‐urea cycle, polyamines, putrescine, reactive oxygen species, stress tolerance, urea cycle

1. Comparative Analysis of the Ornithine‐Urea Cycle in Plants and Animals

The ornithine (Orn)‐urea cycle, also known as the urea cycle, was first described in 1932 by Hans Krebs, a Nobel laureate in 1953, and his assistant Kurt Henseleit in mammals. It was the first metabolic cycle to be elucidated, although the concept of metabolic cycles had already been proposed earlier (Graßhoff and May 2003; Krebs and Henseleit 1932). The urea cycle encompasses five enzymes: carbamoyl phosphate synthase type I (CPS I), Orn transcarbamylase (OTC), argininosuccinate synthase (AS), argininosuccinate lyase (AL), and arginase (ARG; Morris 2002). In mammals, the cycle is primarily located in periportal hepatocytes of the liver, though there is also evidence of low urea cycle activity in enterocytes (Wu 1995). CPS I and OTC are predominantly located in the mitochondrial matrix, whereas AS, AL, and ARG are found in the cytosol (Moedas et al. 2024; Morris 2002).

The enzyme CPS I directly incorporates ammonium (NH4 +) into the urea cycle by producing carbamoyl phosphate, which is then converted to citrulline by OTC (Figure 1). CPS I requires the allosteric activator N‐acetylglutamate, which is synthesized by N‐acetylglutamate synthase and plays a critical role in the regulation of urea biosynthesis in mammals (McCudden and Powers‐Lee 1996; Meijer 1979; Rubio and Grisolía 1981). Utilizing citrulline and aspartate, AS synthesizes argininosuccinate, which is subsequently cleaved by AL to form arginine and fumarate. Subsequently, ARG catalyzes the conversion of Arg to Orn and urea. ARG has two isoenzymes: ARG type I, which is expressed in the liver, and ARG type II, which is found in the kidney (Jenkinson et al. 1996). Cytoplasmic Orn is transported back to the mitochondria to complete the cycle, while urea is excreted (Indiveri et al. 1997; Weiner et al. 2015). Alternatively, Orn can be decarboxylated by Orn decarboxylase (ODC) in the cytoplasm to produce putrescine (Put), the first component of a chain of reactions that generate spermidine (Spd), spermine (Spm), and their acetyl derivatives, catalyzed by the enzymes Spd synthase (SPDS), Spm synthase (SPMS), and Spd/Spm N1‐acetyltransferase (SSAT).

FIGURE 1.

FIGURE 1

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A comparison between the mammalian and plant urea cycle. The urea cycle (in blue), is linked to the polyamine (PA) synthesis (in pink). Carbamoyl phosphate is synthesized from NH4 + by carbamoyl phosphate synthetase I (CPSI) and glutamine by carbamoyl phosphate synthetase II (CPSII). Arginine and ornithine can be used as the precursors of Putrescine which then is then transformed into spermidine and spermine by spermidine synthase (SPDS) and Spermine synthase (SPMS) respectively. An alternative pathway for NO production from Arginine is also represented according to Ricoux et al. (2003) and to Schott et al. (1994). This pathway is still to be confirmed and is depicted with a “?” Pathways not present in plants are represented in red. Enzymes are represented in bold. Glutamine synthetase (GS), Ornithine transcarbamylase (OTC), Argininosuccinate synthase (AS), Argininosuccinate lyase (AL), Arginase (ARG), Urease (URE), Arginine decarboxylase (ADC), Agmatine iminohydrolase (AIH), N‐carbamoyl‐putrescine amidohydrolase (NCPAH), Ornithine decarboxylase (ODC), Copper amino oxidase (CuAO), and Polyamine oxidase (PAO). Modified from Urra et al. (2022).

The efficiency of the urea cycle depends on the metabolic flux of reaction products between the successive enzymes of the cycle (Cheung et al. 1989; Cohen et al. 1987). Additionally, the urea cycle plays a crucial role in the elimination of excess nitrogen (N) as urea in metazoans (Morris 2002) and in the reassimilation of NH4 + in photosynthetic organisms. In lower photosynthetic eukaryotes, genome sequencing suggests that the origin of the urea cycle predates metazoans (Allen et al. 2011; Armbrust et al. 2004; Horák et al. 2020). Phylogenetic analysis indicates that CPS enzymes are present in stramenopiles and haptophytes, in addition to metazoans. CPS genes are thought to have undergone two duplication events, with one lineage giving rise to the type I CPS enzymes (Allen et al. 2011). Consequently, an exosymbiont‐derived Orn‐urea cycle has been identified in diatoms, which relies on a Type I CPS enzyme, similar to the urea cycle in metazoans. However, Type I CPS is absent in green algae and plants. Furthermore, this study revealed that the urea cycle in diatoms functions as a hub for redistributing N and inorganic carbon, contributing significantly to metabolic responses during periods of high N availability. It also serves as a key metabolic pathway for anaplerotic carbon fixation (Allen et al. 2011).

The urea cycle has long been considered incomplete in plants because CPS I, the enzyme responsible for the synthesis of urea from NH4 +, is absent. However, plants possess a functional CPS II enzyme that incorporates glutamine into the cycle instead of NH4 + (Brady et al. 2010; Zhou et al. 2000) (Figure 2, Figure S1). The recent discovery that the urea cycle is down‐regulated in response to NH4 + stress in Medicago truncatula provides evidence that this pathway is also important in plants (Urra et al. 2022). Importantly, the connection between the urea cycle and polyamine (PA) metabolism leads to the synthesis of Put, which plays a vital role in stress protection (Cui et al. 2020).

FIGURE 2.

FIGURE 2

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Schematic representation of CPS evolution including higher plants genes. Modified from Allen et al. (2011). The tree shows the evolution of Carbamoyl phosphate synthase (CPS) within the ornithine‐urea cycle. The ancestral version of CPS present in archaea and bacteria underwent two different duplications and a change of substrate specificity from glutamine to NH4 +. Alignment of different higher plant CPS shows a higher homology to cyanobacterial bacteria and archaea, and ancestral CPS, meaning that plant CPS evolved separately from the tree before the first duplication, 1.500 Myr ago. A more detailed version of this tree can be found as Figure S1.

Beyond this step, plants have all the remaining homologous enzymes involved in the urea cycle found in animals. Similarly to animals, the biosynthesis of Arg in the urea cycle of plants is catalyzed by the sequential action of OTC, AS, and AL (Micallef and Shelp 1989; Slocum 2005). Subsequently, Arg is converted to Orn by the action of ARG, with the concomitant production of urea (Kang and Cho 1990; Winter et al. 2015), which is further degraded by plant urease (URE) to form NH4 + (Polacco and Winkler 1984; Witte 2011; Figure 1). Alternatively, plants can use Arg or Orn to synthesize Put. The decarboxylation of Orn, mediated by ODC, leads directly to Put biosynthesis. From Arg, two different pathways can mediate the biosynthesis of Put (Figure 1). In the first step, agmatine is produced via Arg decarboxylase (ADC). From agmatine, two branches diverge: the first branch produces N‐carbamoyl‐Put, catalyzed by agmatine iminohydrolase, which is then transformed into Put via N‐carbamoyl‐Put amidohydrolase (Figure 1). The second branch produces Put directly from agmatine, catalyzed by ARG/agmatinase (Patel et al. 2017). Finally, Put can be successively aminopropylated to form Spd in a reaction catalyzed by SPDS, and further converted to Spm by the action of SPMS. The donor of aminopropyl groups is decarboxylated S‐adenosylmethionine (dcSAM), which is derived from SAM, a universal methyl donor (Figure 1; Slocum et al. 1984).

The urea cycle and PA synthesis in plants are intricately coordinated pathways that play important roles in plant growth and development. The ability of plants to modulate these pathways, especially the production of PAs, under various environmental stresses such as salinity, drought, and heavy metal exposure, underscores their adaptive strategies to tolerate stress, maintain cellular homeostasis, and enhance survival (Cui et al. 2020; Shao et al. 2022). Understanding the detailed mechanisms and regulatory networks governing the urea cycle and PA synthesis can provide valuable insights for developing crops with improved stress tolerance and overall resilience.

2. Polyamine Production via the Urea Cycle and Its Role in Plant Stress Tolerance

Few studies have addressed the involvement of the urea cycle in plant abiotic stress response. Experimental evidence demonstrates that the regulation of the urea cycle contributes to mercury stress tolerance in the moss Taxiphyllum taxirameum , drought stress in Solanum lycopersicum (tomato) and, more recently, high NH4 + levels in M. truncatula (Hu et al. 2024; Kalamaki et al. 2009; Urra et al. 2022). Under low CO2 conditions (100 ppm), the accumulation of urea cycle intermediates, Orn and citrulline, has been suggested as an alternative sink for photorespiration‐derived N (Blume et al. 2019).

PAs contribute to stress tolerance through their antioxidant property, triggering cell wall modifications, showing acid‐neutralizing ability, and other mechanisms (Ali et al. 2020). Both endogenous synthesis and exogenous application of PAs have been shown to improve plant stress tolerance (Gill and Tuteja 2010). However, the relationship between PAs and stress tolerance is complex. It depends on the plant species, the nature of the PA, its concentration and tissue. In some cases, they have been shown to cause cellular harm through the production of hydrogen peroxide and acrolein during catabolism (Kakehi et al. 2008; Mano 2012; Takano et al. 2012). Despite this duality, elevated PA levels are considered a metabolic hallmark of plant stress (Minocha et al. 2014).

The overexpression of the N‐acetyl‐l‐glutamate synthase (SlNAGS1) gene from tomato in Arabidopsis thaliana resulted in the accumulation of high levels of Orn and increased tolerance to drought and salinity (Kalamaki et al. 2009). The increase in Orn was accompanied by an increase in citrulline and to a lesser extent in Arg content. Also, in Oryza sativa (rice) SlNAGS1 overexpressing lines, the elevated levels of Orn and Arg led to higher production of Spd and Spm, associated with the maintenance of chlorophyll contents and better tolerance to drought (Capell et al. 2004).

Plant responses to water stress exhibit multiple intersections with N metabolism. Cell growth is directly reliant on water influx and is thus rapidly influenced by the limitation in water availability modulating cell wall synthesis (Cosgrove 2024; Hsiao et al. 1997). This early response provokes a marked redistribution of carbohydrates and modulates the carbon‐to‐N ratio at the plant organ level (Gargallo‐Garriga et al. 2014). During drought stress, aerial tissue growth is inhibited while root system activity is either maintained or enhanced, facilitating survival during drought by increasing the root‐to‐shoot ratio (Hsiao and Xu 2000). These changes in carbon allocation across organs necessitate precise regulation of energy metabolism, often associated with reactive oxygen species (ROS) production and the activation of antioxidant pathways (Pinheiro and Chaves 2011). Simultaneously, water stress induces N redistribution, leading to the accumulation of amino acids and nitrogenous compounds, such as PAs (Blázquez 2024). The regulation of cell growth under drought stress significantly impacts protein turnover and proteolytic metabolism, both of which appear to be critical in responding to environmental cues (Eckardt et al. 2024). Among amino acids, proline is notably the most accumulated under drought stress. In addition to its function as an osmoprotectant, proline may also contribute to cellular homeostasis by modulating redox balance, energy status, or acting as a signaling molecule to regulate cell proliferation and gene expression (Szabados and Savouré 2010). Interestingly, proline synthesis appears to associate with an increase in the activity of Δ1‐pyrroline‐5‐carboxylate synthetase and ADC enzymes in the presence of exogenous nitric oxide (NO; Filippou et al. 2013). In this context, the interplay between PA accumulation and proline metabolism may influence the urea cycle, a relationship that might also be shared with other stress conditions, including those induced by NH4 + nutrition.

The positive role of PAs on protecting plants against drought stress has been demonstrated in several plant species, including wheat (Ebeed et al. 2017), maize (Gupta et al. 2012; Hussain et al. 2013), lettuce (Zhu, Dong, et al. 2019; Zhu, Wang, et al. 2019), white clover (Li et al. 2016), trifoliate orange (Fu et al. 2014), and Arabidopsis (Alcázar et al. 2010). In addition, PAs contributed to the protection against drought‐induced oxidative damage in tomato (Sánchez‐Rodríguez et al. 2016). The effect of PAs during drought stress tolerance has been studied with a wide variety of approaches, those including seed priming (Ebeed et al. 2017; Hussain et al. 2013), leaf application (Gupta et al. 2012; Hassan et al. 2020; Zhu, Dong, et al. 2019; Zhu, Wang, et al. 2019), and supplementation to media or nutrient solutions (Fu et al. 2014; Li et al. 2016).

Transgenic Arabidopsis plants overexpressing Arg decarboxylase 2 (ADC2) were shown to accumulate Put in relation to higher drought tolerance (Alcázar et al. 2010). In contrast, the A. thaliana acl5/spms double mutant, which is compromised in Spm and thermospermine biosynthesis, was shown to be hypersensitive to drought stress (Kakehi et al. 2008; Takano et al. 2012). The drought‐sensitive phenotype could be rescued by Spm, but not by Put or Spd pre‐treatments, thus suggesting that the drought‐hypersensitivity was mostly due to Spm deficiency (Yamaguchi et al. 2007). A Put‐to‐Spm canalization was reported in Arabidopsis, which did not increase Spm levels but was linked to drought stress tolerance in the resurrection species Craterostigma plantagineum (Alcázar et al. 2011). The overall data suggest that Spm biosynthesis and oxidation may be relevant in conferring drought tolerance (Figure 1), although the participation of other endogenously synthesized PAs cannot be ruled out. In fact, different effects on stress tolerance are attributed to different PAs, suggesting a distinct specialization in stress signaling (Alcázar et al. 2020; Nandy et al. 2022).

PAs play a critical role in maintaining cellular ROS homeostasis during salt stress (Saha et al. 2015). Salt stress typically reduces the level of Put and Spd, while the Spm level shows species‐specific variation (Santa‐Cruz et al. 1997). PAs also interact with potassium and sodium ions, potentially contributing to the maintenance of cation‐anion balance in leaf tissues (Santa‐Cruz et al. 1997). Recent findings even show that NO works synergistically with PAs to enhance salt tolerance by reducing Na+ toxicity, maintaining ionic homeostasis, and stimulating osmolyte accumulation (Napieraj et al. 2020).

It has also been demonstrated that PAs participate during extreme temperature responses in plants. During cold acclimation, PAs accumulate, with Put showing a particularly distinct accumulation pattern (Alcázar et al. 2011; Cuevas et al. 2009). Spd biosynthesis has been positively correlated with the production of protective compounds such as flavonols, ABA, and antioxidant enzymes, especially in cereals (Gondor et al. 2016). Interestingly, Put appears to influence ABA biosynthesis at the transcriptional level in response to low temperatures, highlighting its modulatory role in hormone homeostasis (Cuevas et al. 2009). The consistency of PA response to cold stress has led to the suggestion that foliar PA levels may serve as biochemical markers for long‐term environmental stress (Minocha et al. 2014).

Under heavy metal stress, PAs are closely linked to increased plant tolerance (Paul et al. 2018). For instance, in sorghum, the application of PAs has been shown to enhance tolerance to Cd, since adding PAs to hydroponic media, Put in particular, significantly influenced Cd accumulation in plant tissues (Kumar et al. 2024). Overall, the urea cycle feeds the metabolic pathway for PA biosynthesis (Figure 1). This is particularly interesting because PAs generally exhibit positive effects on plants under stress conditions, suggesting a potential link between the urea cycle and abiotic stress tolerance.

3. The Role of the Urea Cycle and Polyamines in the Context of Nitrogen Assimilation and Ammonium Stress

Plants mostly take up N in the form of NH4 + and nitrate (NO3 −). The proportion of both N sources has a great impact on cell metabolism and plant performance. Indeed, a high proportion of NH4 + in the medium typically has a detrimental effect on plant growth, a phenomenon known as NH4 + stress. The causes of NH4 + stress are multifactorial and include, among others, pH imbalance, cationic mineral deficiencies, the occurrence of oxidative stress and hormone deregulation, and excessive NH4 + assimilation in the plastids (Esteban et al. 2016; Hachiya et al. 2021).

One of the metabolic responses commonly observed in plants exposed to NH4 + stress is the accumulation of N‐containing metabolites. In this context, several studies have evidenced the accumulation of low C/N ratio metabolites. For instance, Urra et al. (2022) reported the accumulation of intermediates of the urea cycle that would contribute to the channeling of N surplus into: glutamine, Arg, Orn, and Put, which can help plant cells to maintain the endogenous NH4 + content below toxic levels (Ariz et al. 2013; Ueda et al. 2008; Vega‐Mas et al. 2019). This excess of NH4 + also induces the biosynthesis and accumulation of the diamine Put, which serves as a precursor of the triamine Spd and tetramine Spm. The redirection of N towards polyamines is suggested to serve as a cellular strategy to store N under stress conditions and sequester excess NH4 +, thereby mitigating its toxic effects (Majumdar et al. 2016; Moschou et al. 2012; Wuddineh et al. 2018). Thus, these experiments suggest that the modulation of the urea cycle and PA metabolism acts as a protective mechanism against NH4 + stress through connecting C and N metabolism (Gerendás et al. 1997; González‐Hernández et al. 2022; Urra et al. 2022). In particular, Put accumulation has been correlated with the degree of NH4 + stress (Houdusse et al. 2005; Recalde et al. 2021; Urra et al. 2022). However, although the potential role of PAs as N sink cannot be completely discarded, the role of Put in this context seems far beyond simply scavenging NH4 +. Indeed, as previously stated, the accumulation of PAs is common under many abiotic stresses; therefore, suggesting a role for PAs, including Put, as stress‐protective metabolites rather than N‐accumulating molecules. Importantly, in the context of NH4 + stress, to determine whether the accumulation of Put during NH4 + stress was a consequence of the stress or, on the contrary a tolerance response, Urra et al. (2022) supplemented M. truncatula plants with exogenous Put. Their results showed that plants treated with Put exhibited high tolerance when exposed to high NH4 + stress (Urra et al. 2022), which evidences a protective role for Put other than simply acting as a N sink.

In addition, Urra et al. (2022) reported a switch in the functioning of the urea cycle in roots exposed to NH4 + stress, from the ADC‐ to the ODC‐dependent production of Put, suggesting that the ODC enzyme may provide greater plasticity to plants to cope with NH4 + stress. In the same line of evidence, both ADC‐silenced and ODC‐overexpressing tomato mutants showed amelioration of NH4 + toxicity syndrome, associated with Put accumulation (González‐Hernández et al. 2022). Interestingly, three enzymes of the urea cycle can be regulated by NH4 +: urease, agmatine iminohydrolase (AIH) and Put amidohydrolase (NCPAH; Figure 1). In particular, AIH and NCPAH are probable regulators of the diversion of the urea cycle to ODC‐dependent Put formation under high NH4 + supply (Urra et al. 2022).

Furthermore, the accumulation of Put in response to NH4 + stress was also attributed to a reduction in the activity of copper‐containing amine oxidase (CuAO) activity. CuAO activity has been shown to be regulated by feedback mechanisms (Yong et al. 2017), probably through an inhibitory effect of NH4 +. Indeed, in vitro assays supported the existence of a feedback inhibitory mechanism of CuAO by NH4 + (Urra et al. 2022). CuAOs primarily catalyze the oxidation of Put to 4‐aminobutanal while producing NH4 + and H2O2 in peroxisomes and the apoplast (Figure 1). The resulting aldehyde spontaneously cyclizes to form Δ1‐pyrroline, which is further converted into γ‐aminobutyric acid. Although with lower affinity, CuAOs also oxidize Spd and Spm (Ghuge et al. 2015; Planas‐Portell et al. 2013; Qu et al. 2017; Tipping and McPherson 1995; Zarei et al. 2015). PA catabolism contributes to several physiological processes, in part through the release of stress‐related molecules such as GABA and H2O2. Particularly, PA‐derived H2O2 contributes to generate oxidative stress within plant tissues or to activate antioxidative responses and cell wall lignification (Angelini et al. 2008; Gupta et al. 2016; Su et al. 2005).

As stated, one of the causes of NH4 + stress is cation imbalance, because of NH4 + competition with other cations for the same uptake systems (Coleto et al. 2023). Due to their polycationic nature, PAs are fully protonated at physiological pH (Evans and Malmberg 1989). Thus, one of the reasons for PA accumulation under NH4 + nutrition may be the modulation of cell ion balance. Besides, the occurrence of ROS overproduction leading to oxidative stress has been often reported under NH4 + stress. Although unclear, the role of PAs to counteract ROS has been postulated, for instance, through the inhibition of metal autoxidation, a spontaneous, often cyclical process that results in the production of ROS, or by inducing the cell antioxidant machinery (Blázquez 2024).

Glutamine synthetase (GS) is a key enzyme as its reaction to produce glutamine from glutamate and NH4 + is considered the main entrance of N into plant metabolism. This enzyme is responsible for assimilating NH4 + from various sources and recycling N during processes like photorespiration and protein degradation (Bernard and Habash 2009; Miflin and Habash 2002). The production of glutamine from the GS enzymatic activity is also the entrance of N into the urea cycle in plants (Figure 1). GS is encoded by multiple genes, and its regulation is highly complex, occurring at both transcriptional and post‐translational levels. This allows for tissue‐specific expression and dynamic responses to environmental factors (Bernard and Habash 2009). The enzyme is localized in the cytosol and the plastids, and participates in critical physiological processes, including leaf senescence and seed development.

Recent studies have uncovered a novel role for GS‐like enzymes in bacterial PA and monoamine metabolism, suggesting their involvement in stress adaptation and N utilization (Krysenko et al. 2021). Moreover, Silva et al. (2019) demonstrated that GS is regulated by S‐nitrosation in the model legume M. truncatula , with distinct effects on its cytosolic and plastid‐located isoforms. This modification, shown to occur under physiological conditions, highlights the potential role of NO signaling in regulating GS activity at the gate of the urea cycle.

4. Urea Cycle and Polyamine Metabolism Might Be Related to NO Production and Signaling

NO is an essential reactive N species (RNS) in plants and is associated with signaling in several processes such as root organogenesis, mineral uptake, plant defence, stress response, and regulation of auxin levels (Buet et al. 2019; Del Castello et al. 2019; Demecsová and Tamás 2019; Martínez‐Medina et al. 2019; Raya‐González et al. 2019).

The urea cycle and PAs are closely connected to NO regulation. In animals, NO synthase (NOS) is the primary enzymatic source of NO, oxidizing Arg to produce citrulline (Figure 1), both intermediaries in the urea cycle. In rat livers, during endotoxin shocks, inducible NOS (iNOS) are upregulated, while the urea cycle enzyme genes are downregulated (Tabuchi et al. 2000). It is considered that ARG, a key urea cycle enzyme, competes with NOS for Arg, potentially reducing NO production and contributing to vascular dysfunction in various conditions (Durante et al. 2007).

In animals, NOS‐independent production of NO has also been reported, for instance from N‐hydroxy‐l‐Arg (NOHA, Ricoux et al. 2003). NOHA is an intermediate in the first step of NO biosynthesis from l‐Arg by NOS. This step requires oxygen, Ca/calmodulin, and is stimulated by tetrahydrobiopterin (Campos et al. 1995; Stuehr et al. 1991). Interestingly, this reaction can occur without added NADPH, suggesting the presence of an endogenous reductant in NOS that supports Arg oxygenation but not NOHA oxygenation (Campos et al. 1995). NOS can also oxidize NOHA using H2O2, producing citrulline and nitrite/NO3 −, with tetrahydrobiopterin affecting the reaction kinetics (Pufahl et al. 1995). Importantly, in vascular smooth muscle cells lacking NOS activity, NOHA induced dose‐dependent nitrite production, which was inhibited by cytochrome P450 inhibitors but not by NOS inhibitors (Schott et al. 1994). Hence, it is possible that in plants, Arg may also serve as a substrate to produce NOHA. Furthermore, the NOS‐independent NO production from NOHA can be catalyzed by various peroxidases and heme‐containing enzymes. For instance, microperoxidase 8, an engineered hemeprotein, catalyzes NO formation from NOHA in the presence of H2O2 (Ricoux et al. 2003), a step that might as well take place in plants. Hence, Arg may yet be the substrate to NO production and signaling. This NO production pathway may well take place in nearly all compartments, where peroxidases are present, such as the peroxisomes or the cell wall, and be part of specific responses to abiotic and biotic stimuli. This conjecture has to be confirmed yet, and more experimentation is needed to ascertain this aspect.

Numerous studies employing a wide range of methodologies (e.g., Arg‐Cit assay, ozone chemiluminescence‐NO analysis, and spin trapping) have reported Arg‐dependent NO synthesis, often attributing it to NOS activity. This activity has been detected in various species across multiple plant families such as Brassicaceae, Fabaceae, and Asteraceae (Corpas et al. 2009). Notably, studies specifying its subcellular localization have identified NOS activity primarily in chloroplasts and peroxisomes. Interestingly, NO production has also been detected in A. thaliana mitochondria using the DAF‐FM DA (4‐aminomethyl‐2′,7′‐difluorescein diacetate) probe (Guo and Crawford 2005). However, mitochondrial NO production has been attributed to NO2 − reduction in the mitochondrial electron transport chain and was dismissed as non‐“NOS” activity (Modolo et al. 2002).

While NOS‐like activities have been detected in plant extracts, no typical NOS sequence can be found in land plant genomes (Jeandroz et al. 2016). Furthermore, the classical NO/cGMP signaling pathway found in animals is largely absent in plants, with only a few homologues to the NOS enzyme identified in lower algal species (Astier et al. 2019). Instead, land plants appear to have evolved alternative mechanisms for oxidative NO synthesis from amino acids, as the peroxidase‐dependent NO production from oximes described recently (López‐Gómez et al. 2024). These findings suggest that plants have developed unique NO signaling mechanisms, with S‐nitrosation emerging as a ubiquitous NO‐dependent signaling process (Astier et al. 2019).

PAs have been identified as inducers of NO synthesis. Although the specific reaction responsible for this NO production remains unknown, one hypothesis stands that PAs themselves, or a direct derivative, may serve as substrates. Notably, reduced NO biosynthesis has been observed in A. thaliana pao2 knockout mutants (Wimalasekera et al. 2015). This highlights the interconnection between PA catabolism and NO signaling in the modulation of root architecture among other processes (Recalde et al. 2021; Wimalasekera et al. 2011).

Although the relationship between the urea cycle and NO production remains unclear, Arg‐dependent NO synthesis has been reported for long in plant physiology (Corpas and Barroso 2017). Indeed, several studies have shown that common inhibitors of the NOS enzyme also inhibit NO production in plants, indicating a potential overlap in the regulatory mechanisms between plants and other organisms. Tovar‐Mendez et al. (2008) suggested that ARG or ADC may represent alternative pathways for the synthesis of NO in plants from Arg. Indeed, mutation of either ARGs (ARGAH1 or ARGAH2 encoding Arg amidohydrolase‐1 and ‐2, respectively) in A. thaliana resulted in an increased formation of lateral and adventitious roots, effects commonly observed with NO donors and NO production inducers (Flores et al. 2008).

The active role of peroxidases in NO metabolism is now accepted. Besides the role of microperoxidase on NOHA to produce NO, peroxisomes have been reported as the organelle in which l‐Arg‐dependent NOS activity takes place in pea plants. This NO production activity was modulated during senescence, salinity, or Cd stress, while the peroxisomes were shown to contain other NO‐derived molecules such as S‐nitrosoglutathione (Corpas and Barroso 2017). Overall, alternative pathways of NO production from Arg to the animal NOS enzyme exist in plants, which target the urea cycle as a possible subject for the regulation of NO production and signaling in plants.

Given that chloroplasts and peroxisomes are peroxidase‐containing compartments, it is likely that the detected NO production is mediated by organellar‐specific peroxidases, and that the Arg‐NOHA‐NO synthesis pathway is taking place in these organelles (Figure 1). This hypothesis is further supported by multiple reports of peroxidase‐mediated oxidative NO synthesis (Abu‐Soud and Hazen 2000; Del Río et al. 2003; López‐Gómez et al. 2024; Palmerini et al. 2012; Ricoux et al. 2003; van der Vliet et al. 1997). The recently proposed peroxidase‐dependent NO production from oximes is a complex mechanism involving several co‐factors in an intricate enzymatic cycle. Should Arg‐derived NO production be empirically confirmed in plants, Arg would emerge as a central branching point within the urea cycle, regulating two distinct adaptive responses to environmental cues: either PA biosynthesis, promoting autophagy and stress tolerance, or NO production, triggering stress signaling.

PAs have long been recognized as plant growth regulators, since both endogenous biosynthesis and exogenous application influence plant development and stress tolerance. However, the precise mechanisms underpinning PA‐mediated plant responses remain only partially understood (Alcázar et al. 2020; Recalde et al. 2021; Tyagi et al. 2023). More recently, NO has been proposed as a secondary messenger in Put signaling, facilitating the remobilization of phosphorus in root cell walls (Jing et al. 2022) and iron homeostasis (Zhu et al. 2015).

5. Concluding Remarks

The urea cycle functions as a metabolic pathway in higher plants as well as in animals. In contrast to lower photosynthetic eukaryotes, plants acquire N into the cycle by the action of a CPS‐II enzyme which uses glutamine as a substrate. The urea cycle regulates the levels of several important N‐compounds, linking its activity to the upregulation of Put and other PAs, thereby enhancing stress tolerance across various conditions. The stress tolerance increase provoked by PAs seems to be shared across several species, families, and phyla, including plants and mammals, which suggests a possible, yet unidentified, common mechanism.

The studies conducted suggest that NO messenger can be synthesized within the urea cycle, as experiments have shown a triple interplay between the urea cycle, the PA metabolism, and the NO homeostasis. The reactions of peroxidases or cytochrome P450 enzymes on Arg‐derived compounds may be hypothesized as a pathway to NO production in plants, analogously to the animal counterparts. Further studies are required to clarify the role of the urea cycle in PA metabolism and the potential signaling function of PAs in NO release and homeostasis. Overall, a key challenge remains to unravel the precise mechanisms by which Put and other PAs protect plants—and potentially other organisms—against such a wide range of stressors.

Author Contributions

M.U., J.B., and J.F.M. devised the review and wrote the main manuscript. J.B. performed the data analysis and figure design. D.M., E.M.G., and R.A. reviewed the manuscript and added valuable information, input, and references. J.F.M. supervised the whole project.

Supporting information

Figure S1. The phylogenetic tree was inferred from amino acid sequences of carbamoyl phosphate synthetases (CPS) across representative species of higher plants. The sequences were supplemented with those extracted from Allen et al. (2011) and complemented with some others from EMBL Database (Goujon et al., 2010; Madeira et al., 2010). For clarity, in cases where both large and small subunits were present, only the large subunit was included in the analysis.Sequences were aligned using ClustalOmega (Sievers et al., 2011), and the resulting phylogeny was visualized with FigTree v1.4.4 (Rambaut 2010). Abbreviations preceding “CPS” follow the conventions outlined by Allen et al. (2011) and indicate the associated pathway (a, arginine biosynthesis; p, pyrimidine biosynthesis; u, urea cycle) and substrate (g, glutamine; n, ammonium).

PPL-177-e70321-s001.docx (320.3KB, docx)

Acknowledgments

We sincerely thank Dr. Clemence Marchal and Horváth Sára for their support, improvement of the manuscript, and material gathering.

Handling Editor: E. Monte

Funding: This work was supported by grant from MICINN/AEI/10.13039/501100011033/FEDER, UE and the Consolidated Groups program of the Basque Government. J.B. is also a recipient of the Grant “Requalification of the Spanish University System for 2021–2023, Public University of Navarra” fellowship, funded by the European Union‐Next Generation EU/PRTR, Eusko Jaurlaritza, Spain (IT1560‐22). M.U. held a recipient of a predoctoral fellowship from the Government of Navarre, Spain. Open access funding provided by Public University of Navarra.

Data Availability Statement

All data used for this review is publicly available. Data used for the alignments are available upon request.

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Associated Data

This section collects any data citations, data availability statements, or supplementary materials included in this article.

Supplementary Materials

Figure S1. The phylogenetic tree was inferred from amino acid sequences of carbamoyl phosphate synthetases (CPS) across representative species of higher plants. The sequences were supplemented with those extracted from Allen et al. (2011) and complemented with some others from EMBL Database (Goujon et al., 2010; Madeira et al., 2010). For clarity, in cases where both large and small subunits were present, only the large subunit was included in the analysis.Sequences were aligned using ClustalOmega (Sievers et al., 2011), and the resulting phylogeny was visualized with FigTree v1.4.4 (Rambaut 2010). Abbreviations preceding “CPS” follow the conventions outlined by Allen et al. (2011) and indicate the associated pathway (a, arginine biosynthesis; p, pyrimidine biosynthesis; u, urea cycle) and substrate (g, glutamine; n, ammonium).

PPL-177-e70321-s001.docx (320.3KB, docx)

Data Availability Statement

All data used for this review is publicly available. Data used for the alignments are available upon request.

Articles from Physiologia Plantarum are provided here courtesy of Wiley

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