Dressed Up to the Nines: The Interplay of Phytohormones Signaling and Redox Metabolism During Plant Response to Drought

Abstract

Plants must effectively respond to various environmental stimuli to achieve optimal growth. This is especially relevant in the context of climate change, where drought emerges as a major factor globally impacting crops and limiting overall yield potential. Throughout evolution, plants have developed adaptative strategies for environmental stimuli, with plant hormones and reactive oxygen species (ROS) playing essential roles in their development. Hormonal signaling and the maintenance of ROS homeostasis are interconnected, playing indispensable roles in growth, development, and stress responses and orchestrating diverse molecular responses during environmental adversities. Nine principal classes of phytohormones have been categorized: auxins, brassinosteroids, cytokinins, and gibberellins primarily oversee developmental growth regulation, while abscisic acid, ethylene, jasmonic acid, salicylic acid, and strigolactones are the main orchestrators of environmental stress responses. Coordination between phytohormones and transcriptional regulation is crucial for effective plant responses, especially in drought stress. Understanding the interplay of ROS and phytohormones is pivotal for elucidating the molecular mechanisms involved in plant stress responses. This review provides an overview of the intricate relationship between ROS, redox metabolism, and the nine different phytohormones signaling in plants, shedding light on potential strategies for enhancing drought tolerance for sustainable crop production.

1. Introduction

Plants, as sessile organisms, must respond effectively to environmental stimuli to achieve optimal development. Different environmental factors have the potential to inflict stress on plants, limiting their growth. Abiotic stresses, manifested in different ways, are frequently associated with changes in climatic conditions, such as rain, temperature, solar irradiation, and soil quality. Changes in water availability and temperature, leading to hydric and heat stresses, have been related to climate change’s impact [1]. The integration of models on climate change and agricultural production considers that, soon, the abrupt impact of changes in environmental conditions will intensify, representing a greater risk to the productivity of crops. Given plants’ crucial role as a primary food source, this situation could have serious implications for food security in a rapidly growing global population. Moreover, plants are important fuel and fiber sources, contributing significantly to production diversity and ecological sustainability [2,3].

Through evolution, plants have developed adaptive strategies to respond to environmental challenges to ensure their reproductive success. These strategies include a series of morphological, anatomical, and metabolic adjustments to better cope with the environmental conditions. Among these strategies is the crucial importance of reactive oxygen species (ROS) as secondary messengers as well as the signaling mediated by phytohormones, which perform an essential function in plant development and reproduction [4,5]. The integration between hormonal signaling and the management of ROS homeostasis controls diverse types of signaling, especially those involved with stress responses [6].

Many of the signaling pathways in which ROS act as signaling molecules are orchestrated by phytohormones [7], which play an indispensable role in metabolism, development, coordination, stress response, and even in death [8]. Plants synthesize various phytohormones, such as abscisic acid (ABA), ethylene (ET), salicylic acid (SA), jasmonic acid (JA), gibberellins (GA), auxin (AUX), cytokinin (CK), brassinosteroids (BRs), and strigolactones (SL), to orchestrate and regulate various aspects of growth and development. These phytohormones play a fundamental role in a myriad of dynamic yet rigorously regulated molecular responses throughout the plant life cycle. Responding to environmental adversities, these phytohormones play a crucial regulatory role in gene expression machinery, preparing the plant to withstand unforeseen circumstances [9,10,11].

Among the different types of abiotic stress, drought emerges as the main factor that harms crops globally and directly impacts harvests, ultimately restricting total yield potential [12]. The primary mechanisms controlling drought response involve stomatal closure, modulation of root growth and architecture, and the upregulation of anti-stress proteins [13]. As additional strategies that contribute to mitigating the adverse effects of water stress, cells reduce their hydric potential (ψ), leading to the accumulation of specialized solutes. All these multivariate responses are mainly orchestrated by various hormonal regulations, which support cellular plasticity and help plants develop an effective coping strategy against drought [14]. During water shortage, phytohormones regulate plant growth and development, resulting in increased antioxidant enzyme production, secondary metabolites, and heat-shock proteins [15]. Several studies have indicated that communication between ROS and phytohormones signaling is essential to initiate and modulates the response of tolerance of different stresses, including drought. Therefore, precise coordination in the interaction with phytohormones is essential for the appropriate response in plants [7].

To develop comprehensive concepts and strategies for safeguarding plants against the deleterious effects of abiotic stress and meeting the future demands for plant products, an in-depth exploration of molecular-level mechanisms governing plant stress responses is imperative. This review focuses on elucidating the intricate roles played by ROS, redox metabolism, and signaling mediated by the different phytohormones classes—abscisic acid, ethylene, salicylic acid, jasmonic acid, gibberellins, auxin, cytokinin, brassinosteroids, and strigolactones—in the context of drought stress. The objective is to provide a comprehensive overview of the existing knowledge about how these signaling cascades and redox metabolic processes synergize, leading to robust responses to water stress in plants.

2. Abscisic Acid

ABA is a phytohormone with a central role in the response to water and osmotic stress. ABA orchestrates a diverse range of physiological processes, from seed germination to stomatal regulation. Simultaneously, it controls plant growth and development by regulating the synthesis of protective metabolites. These metabolites enable plants to combat the detrimental effects of stressful environments, which is why ABA is often referred to as the “universal stress hormone”.

2.1. Abscisic Acid Metabolism

In the ABA biosynthesis pathway, the enzyme 9-cis-epoxycarotenoid dioxygenase (NCED; EC 1.13.11.51) acts as a bottleneck in the conversion of 9-neoxanthins to xanthin. This enzymatic process influences ABA levels during drought response. The subsequent enzymatic cleavage of ABA into 80-OH-ABA, catalyzed by ABA 8-hydroxylase (CYP707A; EC 1.14.14.137), represents an inactivation mechanism, resulting in dehydrophaseic acid (DPA) and phaseic acid (PA) as the main degradation products [16]. ABA, primarily produced in leaf vascular tissues in response to water deficit stress, is transported to guard cells, where it triggers stomatal closure. It is believed that ABA is transported via passive diffusion from a low- to a high-pH environment due to its nature as a weak acid, without the need for ABA transporters [10]. However, ABA can also be transported by specific ABA transporters, particularly members of the ABC subfamily G [17] and the NPF (Nitrate transporter 1/peptide transporter family) [18].

2.2. Abscisic Acid Signaling

Under basal conditions, plants produce ABA at low concentrations [19], and the kinase activity of SnRK2 (sucrose nonfermenting 1 (SNF1)-related protein kinase 2) is inhibited by the phosphatase PP2C. In arabidopsis (Arabidopsis thaliana), PP2Cs from clade “A”, such as HAI1 (HIGHLY ABA-INDUCED 1), HAI2, HAI3, AHG1 (ABA-HYPERSENSITIVE GERMINATION 1), AHG1, RDO5 (REDUCED DORMANCY 5), ABI1 (ABA-INSENSITIVE 1), ABI2, HAB1 (HYPERSENSITIVE TO ABA1), and HAB2, have been identified as important regulators of ABA signaling [20,21]. The increase in ABA concentration triggers ABA signaling, which involves the recognition of ABA by PYR/PYL/RCARs receptors, leading to the inactivation of PP2C phosphatase activity (Figure 1) [22,23].

The PYR/PYL/RCAR proteins are encoded by various genes, acting as essential receptors for the ABA response. The autoactivation of SnRK2/OST1 is triggered when the protein dissociates from the PP2C phosphatases. In arabidopsis, the isoforms SnRK2.2, SnRK2.3, and SnRK2.6 are primarily involved in the ABA response and function by phosphorylating different targets [21,24].

The ABA signaling cascade involves several transcription factor branches, and the cis-acting element known as the ABA-responsive element (ABRE) is present in the promoters of genes induced by drought [25]. These ABREs function as binding sites for leucine zipper transcription factors, such as ABRE binding protein/ABRE binding factors (AREBs/ABFs) and ABI3 (ABA-INSENSITIVE 3), ABI4, and ABI5 [25]. During ABA signaling, ABA-dependent SnRK2 protein kinases phosphorylate and directly activate the AREB/ABF and ABI transcription factors [24,26] (Figure 1).

ABA-responsive elements contribute significantly to the complexities of ABA signaling [27]. SnRK2 activation triggers the induction of molecular and physiological responses to ABA, such as the modulation of photosynthetic activity, root growth, and germination [21]. In addition to these direct effects, ABA exerts regulatory control over a series of genes associated with drought responses, substantially contributing to drought tolerance development in plants [15,28]. This finding, coupled with the identification of numerous classic ABA-insensitive mutations located in transcriptional regulators, strongly suggests a central role in gene regulation, conferring resistance to abiotic stress [29].

2.3. Abscisic Acid and ROS on Drought Response

During drought response, ABA also plays an important role in regulating ROS production, as hydrogen peroxide is generated mainly by the interaction and phosphorylation of NADPH oxidases (known as respiratory burst oxidase homologs, (RBOHD and RBOHF)) via SnRK2 [30,31] (Figure 1). Furthermore, SnRK2-dependent hydrogen peroxide production can act as a signal for further release of active SnRK2 through PP2C inactivation [32]. Consequently, ABA and ROS act in a positive feedback cycle, resulting in elevated levels of ROS/ABA that regulate gene expression and facilitate cellular responses to stress [33].

The ABA-mediated ROS production controls stomatal opening by activating Ca²⁺ channels in guard cells, triggering an increase in the concentration of cytoplasmic Ca²⁺ ([Ca²⁺]cyt) that then activates SLAC1 (slow anion channel 1) via CPK/CDPK (calcium-dependent protein kinase) or CBL (calcinuerin B-like) [17]. The ABA-induced hydrogen peroxide activates guard cells’ GHR1 membrane receptor protein kinase (GUARD CELL HYDROGEN PEROXIDE-RESISTANT 1), which interacts and activates SLAC1 [34]. Both CPK and GHR1 are also able to activate SLAH3 (SLAC1 homolog 3), promoting the Cl⁻ efflux [35]. This effect is accompanied by the regulation of other membrane ion transporters by SnRK2, such as the activation of outward shaker K⁺ channels GORK, KUP6, and KUP8 and the inhibition of both inward K⁺ channels KAT1/KAT2 and the aluminum-activated malate transporter ALMT12/QUAC1 channel (quick anion channel 1) [17] (Figure 1). Anion and K⁺ effluxes cause membrane depolarization and reduction in guard cell turgor, ultimately leading to reduced stomatal aperture size and minimized water loss through transpiration [33,36]. This mechanism is directly related to increased hydrogen peroxide levels: the double mutant atrbohd/atrbohf, which generates lower ABA-induced ROS, showed impaired ABA-induced Ca²⁺ channels and stomatal closure [30]. This phenotype is rescued by exogenous hydrogen peroxide, indicating that ROS serves as a secondary messenger in ABA signaling of stomatal response [37]. ABA-induced hydrogen peroxide also activates NIA1 and NIA2 (nitrate reductase 1 and 2; EC 1.7.1.1), leading to NO (nitric oxide) synthesis, which is also critical for ABA-induced stomatal closure [38]. NO negatively regulates abscisic acid signaling in guard cells by inhibition of ABA receptor PYR/PYL/RCAR [39] and SnRK2 [40] (Figure 1). NO also reduces ABA content by the induction of the expression of ABA catabolic gene CYP707A and the inhibition of ABA biosynthesis-related gene NCED1 [41]. Indeed, enhanced sensitivity to ABA during development and in responses to stress is observed in the NO-deficient arabidopsis triple mutant nia1 nia2 noa1-2 [42]. ROS signaling also plays a significant role in water stress tolerance through ABA-induced transcription factors. In arabidopsis, RRTF1 (redox-responsive transcription factor 1), belonging to the ERF/AP2 (ethylene response factor 1; APETALA 2) family, is a key component of the central redox signaling network. This network is activated by both ABA and ROS in response to various stresses, including drought. Elevated levels of RRTF1 lead to additional ROS accumulation, playing an important role in initiating subsequent acclimation responses [43,44].

During water stress, ABA also regulates root tissue patterning. In the endoderm, ABA signaling promotes xylem differentiation by inducing the expression of microRNAs miR165 and miR166, both essential regulators of vascular development [45]. In addition, ABA acts directly in xylem cells, activating the expression of VND (vascular-related NAC domain) transcription factors that promote xylem differentiation [46].

3. Ethylene

Ethylene is a gaseous phytohormone, which governs various physiological processes and responses to environmental stimuli, regulating leaf abscission, flowering, fruit ripening, senescence, seed germination, cell division, tissue differentiation, sex determination, adventitious root growth, starch accumulation epinasty, stomatal closure, etc. [47,48,49,50,51,52,53,54,55]. Moreover, it acts as a triple-response growth regulator, influencing shoot elongation, stem thickening, and horizontal growth habit [47], and it serves as a stress hormone, mediating plant reactions to biotic and abiotic stresses like pathogen attacks, salinity, drought, hypoxia, cold, and heat [56,57,58,59,60,61].

3.1. Ethylene Metabolism

The ethylene biosynthesis pathway starts with the amino acid methionine, which is converted to S-adenosyl-L-methionine (SAM) by S-adenosylmethionine synthetase (SAMS; EC 2.5.1.6) in the Yang cycle [62,63]. SAM is then converted into 1-aminocyclopropane-1-carboxylic acid (ACC) and 5-methylthioadenosine (MTA) by ACC synthase (ACS, EC 4.4.1.14) [63]. While MTA is recycled through the Yang cycle reactions back to methionine [64], ACC is oxygenated by ACC oxidase (ACO, EC 1.14.17.4; also named ethylene forming enzyme—EFE) to produce ethylene [65]. Ethylene biosynthesis is regulated mainly by the transcriptional and post-translational control of ACS and ACO steps by different internal and external stimuli [66].

3.2. Ethylene Signaling

The classic ethylene signal transduction is triggered by gas interaction with endoplasmic reticulum (ER) membrane receptors, including ETHYLENE RESPONSE 1 (ETR1), ETR2, ETHYLENE-INSENSITIVE 4 (EIN4), ETHYLENE RESPONSE SENSOR 1 (ERS1), and ERS2 [67], and the modulation of CTR1 (CONSTITUTIVE TRIPLE RESPONSE 1; EC 2.7.11.1) activity to regulate the expression of several genes (Figure 2).

The ethylene receptors are active in the absence of ethylene and constitutively activate the kinase activity of CTR1 [68], which is a Raf-like kinase able to phosphorylate the C-terminal domain of EIN2 (ETHYLENE-INSENSITIVE 2), which is also present in the ER membrane, leading its degradation by proteolysis. The EIN2 C-terminal domain previously phosphorylated by CTR1 interacts with ETP1 and ETP2 (EIN2 targeting proteins), which have F-box domains and are targeted for ubiquitination and degradation by the 26S proteasome [69]. The receptors activity can be positively regulated by complex formation with RTE1 (REVERSION TO ETHYLENE SENSITIVITY 1) and ARGOS (auxin-regulated gene involved in organ size) proteins, considered as negative regulators of ethylene sensitivity [70,71,72,73,74]. The active ethylene receptor biogenesis and the ethylene sensibility are also regulated by the availability of the copper cofactor, which is provided by the copper transporter RESPONSIVE-TO-ANTAGONIST 1 (RAN1), located in the ER membrane [75,76]. Upon binding with ethylene, the ethylene receptors undergo conformational changes, impairing the activation of CTR1. Consequently, instead of being phosphorylated, the EIN2 C-terminal domain is cleaved from the ER membrane by a protease enzyme and transported to the nucleus, thereby initiating ethylene signaling (Figure 2).

In the nucleus, EIN2 stimulates EIN3/EIL (ETHYLENE-INSENSITIVE 3; EIN3-like) transcription factors activity by repressing EBF (EIN3 binding F-box protein), which constitutively binds to EIN3/EIL, targeting it to ubiquitination and degradation [77]. Thus, under ethylene response, EIN3/EIL regulates the expression of ethylene-response genes by binding to the specific cis-acting AGCCGCC motifs (GCC box) and dehydration-responsive element (DRE) [78]. This pathway includes the ERF/AP2 transcription factor superfamily [79,80,81,82] that regulates the molecular response to different stresses, such as drought, salt, heat, and pathogen attack [78]. Interestingly, the EBF mRNA levels can be regulated by the exoribonuclease 5′-3′ EIN5 (ETHYLENE-INSENSITIVE 5, EC 3.1.1.3), another positive regulator of ethylene response, which promotes EBF mRNA decrease, thereby contributing to prevent EIN3/EIL proteolysis [83].

In addition to the canonical CTR1-dependent pathway, the ethylene signal also operates through noncanonical pathways. The EIN3/EIL transcription factors, for example, can be activated by alternative pathways, which involves a phosphorylation cascade of kinase proteins: MKK4-5-9 → MPK3-6 [83,84,85]. Once activated, MPK3-6 phosphorylates EIN3/EIL, preventing their interaction with the F-box protein EBF (EIN3 binding F-box protein). As a result, these factors accumulate in the nucleus and interact with target gene promoters, initiating various ethylene responses [85]. (Figure 2).

The noncanonical pathways also allow the ethylene receptors to interact with components of the cytokinin signaling [72,86,87,88,89]. The ETR1, a canonical histidine kinase, can directly interact with the histidine-containing phosphotransferase protein 1 (AHP1 in arabidopsis) through its C-terminus domain [87,90]. AHP1 is also phosphorylated by the cytokinin receptors by a phosphor-relay pathway and transfers the phosphate group to the response regulator proteins (ARR family in arabidopsis) that function as transcription factors during the cytokinin response [91]. Previous work demonstrated that the arabidopsis arr1 mutant showed reduced ethylene sensitivity in the regulation of root apical meristem size [92]. Indeed, ETR1 histidine kinase activity integrates ethylene and cytokinin signaling, inhibiting root growth [89]; nevertheless, this mechanism is not fully understood.

3.3. Ethylene and ROS on Drought Response

Under stress conditions, ethylene can elicit various responses, including the upregulation of antioxidative enzymes to scavenge ROS and alleviate oxidative stress as well as adjustments in plant physiology. During drought and salt stress, ethylene is known to decrease stomatal aperture, stimulate the growth of adventitious roots. and promote leaf senescence. These adjustments have a direct impact on plant stress response: (i) the stomatal closure stops water loss, (ii) the increased adventitious roots aid water absorption, and (iii) senescence may lead to the redistribution of nutrients in the sink organs [93]. Furthermore, under drought conditions, ethylene has been demonstrated to induce osmolyte biosynthesis [94] and leaf abscission, thereby reducing water loss [48,95]. The drought response mediated by ethylene can be highly regulated by hydrogen peroxide production via RBOHF (respiratory burst oxidase homolog protein F), which is inhibited by CTR1 [96]. In the presence of ethylene, CTR1 is deactivated, thus triggering the RBOHF-dependent hydrogen peroxide production in a mechanism mediated by Gα protein [97]. The increased hydrogen peroxide in guard cells also contributes to the cleavage and nuclear localization of EIN2 through the activation of MKK1/3–MPK3/6 signaling cascade [96]. This pathway induces the expression of nitrate reductase NIA1, which is also activated by hydrogen peroxide, contributing to the increase in NO synthesis. The NO, in turn, activates SLAC1 through both H₂S-dependent and -independent pathways, leading to stomal closure [98]. Indeed, ethylene-induced NO synthesis activates L-cysteine desulfhydrase (DES1; EC 4.4.1.28), inducing H₂S production and stomata closure [99]. Despite this evidence, ethylene is also able to inhibit the ABA-induced stomatal closure or to reduce stomatal sensitivity to stresses in different species [98,100]. These data suggest that while ABA is considered to act as the key regulator of stomatal closure under abiotic stress, ethylene can act as either a positive or negative regulator [101]. Thus, the mechanism by which ethylene regulates stomatal behavior during drought stress as well as the ethylene–ABA interaction pathway is complex and seem to be strongly dependent on their endogenous levels. This dual role during drought stress has led to ethylene and ABA being labeled “Foes or Friends” [101].

The ability of ethylene to either induce or inhibit stomatal closure during drought stress can be partially elucidated by the roles of its receptors. Among the five ethylene receptors described, only ETR1, EIN4, and ERS1 are involved in ethylene-induced stomatal closure, whereas ETR2 and ERS2 receptors may be involved in other ethylene-induced processes [97,102].

4. Salicylic Acid

Salicylic acid is a key signal molecule in regulating the activation of local and systemic defense responses against pathogens infections through the induction of defense genes [103,104,105]. Despite this, salicylic acid was first reported as a signaling molecule in plants, regulating the thermogenesis phenomenon in reproductive organs of gymnosperms and angiosperms [106,107]. Additionally, salicylic acid plays a regulatory role in abiotic stresses, like heat, cold, salinity, and drought [108,109,110], and influences plant growth and development by regulating various processes, including photosynthesis, respiration, vegetative growth, seed germination, flowering, and senescence [111,112,113].

4.1. Salicylic Acid Metabolism

Salicylic acid is a phenolic compound synthetized in plants via the isochorismate synthase (ICS, EC 5.4.4.2) or the phenylalanine ammonia-lyase (PAL, EC 4.3.1.5) pathways. Pathogen-induced salicylic acid is mainly synthesized via the ICS pathway in chloroplasts, which is mediated by three classes of enzymes: (i) ICS1, also known as SID2 (SALICYLIC ACID INDUCTION DEFICIENT 2) [114]; (ii) AtGH3.12 (also called AtPBS3 or avrPphB SUSCEPTIBLE 3), a family of cytosolic acyl adenylase proteins responsible for the conjugation of glutamate to isochorismate (IC) to form IC-glutamate; and (iii) EPS1 (ENHANCED PSEUDOMONAS SUSCEPTIBILITY 1), a BAHD acyltraferase-like protein responsible for the conversion of IC-glutamate to salicylic acid. This last step can also occur spontaneously [115,116,117]. Although the ICS pathway is fully elucidated, all three biosynthetic genes in this pathway, ICS1, GH3.12, and EPS1, are only found in the Brassicaceae family [116,118,119], while the PAL pathway is part of the phenylpropanoid metabolism and appears to be ubiquitous [117].

4.2. Salicylic Acid Signaling

Given the diverse roles of salicylic acid in environmental stress responses, it is likely that plants possess multiple salicylic acid receptors, and the protein NPR1 (NONEXPRESSOR OF PATHOGENESIS-RELATED GENES 1) is a redox-sensitive master regulator of salicylic acid signaling [120]. In the absence of salicylic acid, NPR1 is found oligomerized in cytosol. During stress response, when salicylic acid synthesis is induced, there is a change in the redox state at the cytoplasm, leading to the monomerization of NPR1 proteins. In its monomeric form, NPR1 is translocated to the nucleus, where it activates the expression of salicylic acid-responsive genes, including PR genes [121] and those involved in the induction of the hypersensitive response (HR) [103,122] (Figure 3).

The redox modification of cysteine residues in the protein–protein interaction and the cryptic trans-coactivation domains of NPR1 is essential for its monomerization [120,123,124]. Indeed, in arabidopsis and rice (Oryza sativa), mutations in conserved Cys82 and Cys216 of NPR1 resulted in this constitutive monomerization and nuclear localization of NPR1, resulting in the activation of defense genes expression [125]. Salicylic acid-induced AtNPR1 monomerization and nuclear import is catalyzed by thioredoxins AtTRXh3 and AtTRXh5, while in rice and tobacco, these processes are mediated by the orthologs OsTRXh2 and NtTRX1 [126,127]. In the absence of salicylic acid, NPR1 is also subject to degradation via the 26S proteasome [128], indicating that the regulation of NPR1 protein level is essential to salicylic acid response. NPR1 degradation is dependent on its interaction with NPR3 and NPR4 proteins, which have low and high affinity to salicylic acid, respectively [129]. While salicylic acid disrupts NPR1–NPR4 interaction, it facilitates NPR1–NPR3 interaction, creating an optimal NPR1 concentration gradient to regulate the salicylic acid response. In the absence or low concentration of salicylic acid, NPR4 mediates NPR1 degradation, whereas excessive salicylic acid levels prevent NPR1 accumulation by enhancing its interaction with NPR3 [130] (Figure 3). Thus, NPR1-mediated signaling is active only at intermediate salicylic acid levels. This model aligns with observations that NPR1 accumulates in regions adjacent to the pathogen infection sites, whereas at the infection site itself, the salicylic acid levels are even higher.

The salicylic acid response is mainly mediated by the upregulation of pathogenesis-related (PR) genes. The PR proteins have ~5–43 kDa and are thermostable, protease-resistant, and ubiquitously expressed, representing up to 10% of the total protein in leaves [131,132]. The PR proteins form at least 17 families, with distinct structure and function triggered by different pathogens [133,134]. The response to fungal attack, for example, involves PR1 (CAP-domain proteins), PR2 (β-1,3-glucanases), PR3 (chitinases), PR4 (Barwin-domain proteins), PR5 (thaumatin-like), PR12 (antimicrobial defensins), and PR17 (NtPRp27-like proteins) [132,134,135,136]. Additionally, salicylic acid positively regulates the biosynthesis of different defense molecules, such as flavonoid and phytoalexins [137]. This regulation is mainly mediated by NPR1 that regulates the transcription of salicylic acid-responsive genes mainly through its interactions with TGA (TGACG-binding) transcription factors, a subclass of the basic leucine zipper (bZIP) transcription factor family [138] (Figure 3). Besides the functional activation of transcription factors through complex formation, NPR1 also controls the expression of transcription factors genes, essential for salicylic acid-mediated transcriptional reprogramming, such as WRKY [139,140].

Although the salicylic acid signaling has not been fully characterized, it is known that salicylic acid regulates ROS production through mitochondrial respiration [141] and various enzymes related to ROS metabolism, such as CAT [142] and peroxidases (POD) [143]. At high concentrations, salicylic acid inhibits mitochondrial succinate dehydrogenase (SDH, EC: 1.3.5.1) and the respiration in tobacco (Nicotiana tabacum) cells [143] and in arabidopsis isolated mitochondria [144,145,146] (Figure 3). Conversely, at lower concentrations, salicylic acid induces SDH activity and mitochondrial oxygen consumption [141,145]. These findings suggest that salicylic acid signaling regulates mitochondrial respiration, leading to differential patterns of respiration and ROS production during the plant stress response. Indeed, a point mutation in SDH1-1 subunit (disrupted stress response 1- dsr1), which lowers SDH activity, results in decreased mitochondrial ROS production and impairs the stress response mediated by salicylic acid [147].

4.3. Salicylic Acid and ROS on Drought Response

Salicylic acid is also involved in abiotic stress tolerance [109,148], as it is increased under drought stress [149]. Salicylic acid induces stomatal closure [150,151] and decreases CO₂ assimilation [152], inhibiting water loss and photosynthesis. Indeed, drought-induced ROS generation mediates salicylic acid-induced stomatal closure [130,153]. While plasma membrane NAD(P)H oxidases are the leading producers of ROS in guard cells induced by ABA signaling [30], salicyl-hydroxamic acid (SHAM)-sensitive peroxidases (PRX) are major ROS sources in guard cells induced by salicylic acid signaling [154]. The apoplastic PRX33 and PRX34, which are preferentially expressed in guard cells [155], are deeply involved in hydrogen peroxide production in response to various stimuli (including fungi, bacteria, and flg22), leading to stomatal closure [156,157,158,159,160]. Thus, the ROS generated by salicylic acid integrate into the ABA-induced stomatal closure pathway, activating calcium channels and inducing calcium influx [161] (Figure 3). Notably, salicylic acid-mediated ROS production and stomatal closure are not affected by NADPH oxidase inhibitor DPI (diphenyleneiodonium) or in the rbohd/rbohf double mutant [151].

Salicylic acid also regulates stomatal closure by increasing ethylene synthesis via induction of ACS2, ACS6, and ACS11 [162,163]. Indeed, ethylene biosynthetic inhibitors and mutations in ethylene-signaling genes (ETR1, EIN2, and EIN3) can inhibit stomatal closure induced by salicylic acid, which further suggests ethylene function downstream of salicylic acid to induce stomatal closure during drought [163]. Considering all these data together, ethylene and salicylic acid signaling may complement each other to induce stomatal closure during drought when ABA signaling is not sufficient [98].

5. Jasmonic Acid

Jasmonic acid acts both as a growth regulator and as a crucial signaling molecule in plant defense mechanisms [164]. Jasmonic acid orchestrates plant development and responds to a spectrum of biotic stresses, including physical injury, pathogen attacks, and insect predation, as well as numerous abiotic stresses, like low temperature, drought, excessive salinity, and UV radiation [165,166]. However, the intricate jasmonic acid-mediated signaling network, vital for safeguarding against external threats, can sometimes inhibit plant growth and development due to its over-activation by various regulatory factors [167].

5.1. Jasmonic Acid Metabolism

In response to developmental and environmental cues, plants rapidly produce a substantial increase in jasmonic acid, with is synthesized from α-linolenic acid through a series of reactions that occur sequentially in plastids and peroxisomes. Jasmonic acid–isoleucine conjugate (JA-Ile), the most biologically active component in jasmonate cascade, is synthesized by an ATP-dependent jasmonate–amino acid amidase encoded by JAR1 (JASMONATE RESISTANT 1) [168,169,170,171]. On the other hand, the ω-oxidation pathway, involving members of CYP94B3, mediates the turnover and inactivation of JA-Ile, which is converted to 12-hydroxy-JA-Ile (12-OH-JA-Ile) [172].

5.2. Jasmonic Acid Signaling

Jasmonic acid and JA-Ile subcellular distribution is controlled by JAT (JASMONATE TRANSPORTER), which is an ABC transporter (ABCG16 in arabidopsis) with an unexpected dual location at both the plasma membrane and nuclear envelope [173]. Jasmonic acid signaling is initiated by the hormone being perceived by the COI1 (CORONATINE-INSENSITIVE 1) nuclear receptor, which, upon jasmonate binding, interacts with the repressor JAZ (jasmonate-ZIM-domain) protein. In this context, COI1 binds to other proteins to form the Skp1/Cullin/F-box (SCF^COI1) ubiquitin ligase complex, which ubiquitinates the JAZ protein, targeting it for degradation (Figure 4) [174,175]. In the absence of jasmonic acid, JAZ proteins recruit the protein TPL (TOPLESS) and the adaptor protein NINJA (Novel INteractor of JAZ) to form an effective transcriptional repressor complex [176] that inhibits various transcription factors, including MYC2 (MYELOCYTOMATOSIS 2). The MYC2 proteins are the main transcription factors in jasmonic acid signaling, exerting significant influence on the transcriptional reprogramming of several stress-responsive genes [167]. MED25 (Mediator 25), also known as PFT1 (PHYTOCHROME AND FLOWERING TIME 1), connects MYC2 with RNA polymerase II, thereby modulating gene transcription [177,178,179]. The JAZ–NINJA–TPL complex also recruits histone deacetylases HDA6 and HDA19, preventing the activation of jasmonate-responsive genes [177,180,181,182] (Figure 4).

Jasmonic acid plays an important role in plant biotic stress responses, triggered mainly by insects or pathogens. Jasmonic acid enhances the production of toxic proteins such as the defensive enzyme PPO (polyphenol oxidase), which affects the digestive physiology of herbivories, thereby protecting plants from herbivory [183,184]. Additionally, many secondary metabolites, such as tannins, phenols, flavonoids, and lignin, are induced by jasmonic acid and contribute to the resistance to biotic stress [185,186]. Through MYC2 activation, jasmonic acid regulates the accumulation of ROS through RBOHD and RBOHF [187], which act as a downstream signal following JAZ resulting in systemin-induced defense genes in response to wounding [188]. Like wounding, jasmonic acid also leads to a strong NO burst [189], which is involved in some jasmonic acid-mediated defense responses, inhibiting hydrogen peroxide accumulation [190,191].

Although salicylic acid regulates plant defense responses against some pathogens, salicylic acid and jasmonic acid signaling are antagonists [192]. While jasmonic acid-mediated defense is related to necrotrophy, the salicylic acid-mediated defense is related to hemibiotrophs [117]. The COI1 receptor regulates jasmonic acid-mediated inhibition of the salicylic acid pathway [193]. In a similar way, NPR1 not only controls salicylic acid signaling but antagonizes jasmonic acid signaling [194]. Jasmonic acid modulates the interaction between MYC2 and three NAC transcription factor genes (ANAC019, ANAC055, and ANAC072) to block the accumulation of salicylic acid by inhibiting the expression of genes involved in salicylic acid biosynthesis [195]. Additionally, MPK4 serves as a positive regulator of GRX480 in the salicylic acid signaling but negatively regulates MYC2 in the jasmonic acid signaling [196].

5.3. Jasmonic Acid and ROS on Drought Response

The activation of jasmonic acid-mediated signaling also contributes to plant tolerance to water stress [27]. Arabidopsis mutants of jasmonic acid signaling coi1-2, jaz1, and myc2-2 exhibit drought sensitivity, indicating the importance of signal transduction mediated by jasmonic acid in drought response [197]. Additionally, during the response to drought in rice, OsJAZ proteins interact with the transcription factor OsbHLH148 and induce the expression of the DREB (DEHYDRATION-RESPONSIVE ELEMENT-BINDING) protein, contributing to drought tolerance [198]. During drought stress, jasmonic acid contributes to reducing stomata apertures [199]. Indeed, external application of jasmonic acid induces stomatal closure, triggering a robust response of plants to water stress [200]. This mechanism occurs with or without ABA contribution and is related to ROS production via RBOHD and RBOHF enzymes [199]. 12-OPDA (12-oxo-phytodienoic acid), one of the precursors of jasmonic acid, is also able to induce stomatal closure. Dry conditions prevent the conversion of 12-OPDAs into jasmonic acid, leading to 12-OPDA accumulation, which stimulates stomatal closure, either in conjunction with or independently of ABA, preventing water loss [15,201,202].

Jasmonic acid also boosts the activity of antioxidant enzymes such as superoxide dismutase (SOD), catalase (CAT), ascorbate peroxidase (APX), monodehydroascorbate reductase (MDHAR), dehydroascorbate reductase (DHAR), glutathione peroxidase (GPX), glutathione reductase (GR), glutathione S transferase (GST), and G-POD (guaiacol peroxidase) [203,204], providing protection against oxidative damage and increasing stress tolerance [205]. During drought stress, jasmonic acid-induced NO regulates the AsA-GSH cycle activity, contributing to the maintenance of AsA/DHA and GSH/GSSG ratios, protecting plant cells against oxidative damage [206].

6. Gibberellins

Gibberellins are tetracyclic diterpenoid carboxylic acids and play a crucial role in plant growth, regulating crucial aspects throughout both the vegetative and reproductive phases of a plant’s life cycle, such as seed germination, immature and mature stages of plant development, and flowering time. Additionally, gibberellins regulate the response to different abiotic stresses, including drought [15].

6.1. Gibberellins Metabolism

Gibberellins are synthesized from geranylgeranyl diphosphate (GGDP) through the terpenes pathway. Among the 136 gibberellins structurally characterized, a few have been shown to be biologically active, such as GA₁, GA₃, GA₄, and GA₇ [207]. The non-biologically active GA₁₂ is the first gibberellin in this biosynthesis route, and through the activities of different GA-oxidases, such as GA-20ox and GA-3ox (EC 1.14.11 and EC 1.14.11.15), it is converted into active gibberellins as well as other inactive intermediaries [208]. Gibberellin catabolism and deactivation occur mainly by oxidation reactions carried out by GA-13ox (EC 1.14.-.-) [209] and GA-2ox (EC 1.14.11.13) [210]. Alternatively, the formation of methyl esters by GAMT1 (gibberellin methyl transferase-1) is also able to inactivate gibberellins [211]. Gibberellins can be transported across the plasmatic membrane through transporters from SWEET (SUGAR WILL EVENTUALLY BE EXPORTED TRANSPORTER) [212] and NPF [213] (Figure 5).

6.2. Gibberellins Signaling

In the absence of gibberellin, the DELLA repressors proteins, such as SLR1 (SLENDER RICE 1) in rice and GAI (GIBBERELLIC ACID-INSENSITIVE), RGA (REPRESSOR-OF-ga1-3), RGL1 (RGA-LIKE 1), RGL2, and RGL3 in arabidopsis, act as primary suppressors of gibberellin hormonal responses based on the specific protein interaction partner [214]. DELLA proteins are recruited to target promoters by binding to transcription factors via the LHR1 subdomain [215]. Additionally, the DELLA–histone 2A interaction, via the PFYRE subdomain, is crucial for stabilizing the TF–DELLA–histone 2A complex at the target chromatin, mediating global transcriptional regulation [216].

Gibberellin signaling is initiated by its perception by the nuclear GID1b/c receptor (GIBBERELLIN-INSENSITIVE DWARF 1), leading to the ubiquitination of DELLA repressor protein through SCF (S-PHASE KINASE-ASSOCIATED PROTEIN 1/CULLIN 1/F-BOX)-type E3-ubiquitin ligase) complexes, resulting in its degradation via 26S proteasome and culminating in phytohormone response activation (Figure 5). In rice, DELLA ubiquitination is provided by SCFGID2 (SCF complex associated with the F-box protein subunit GID2) [217,218], while in arabidopsis, DELLA is ubiquitinated by SCFSLY1 or SCFSNE (SCF complexes associated with the F-box protein subunit SLEEPY1 or SNEEZY) [219,220,221,222,223]. Although DELLA proteins are typically inactivated by protein degradation, the gibberellin-induced interaction with the GID1 receptors is sufficient to inhibit DELLA activity when DELLA ubiquitylation and proteasomal degradation are blocked [214]. This mechanism is confirmed in rice gid2 and arabidopsis sly1 mutants, which show a comparatively mild phenotype and accumulate very high levels of DELLA proteins [224,225].

DELLAs interact and inhibit different transcription factors related to different aspects of plant development, such as the phytochrome-interacting factors PIF3 and PIF4, involved in photomorphogenesis [226,227]; ALC (ALCATRAZ) and SPT (SPATULA), repressors of cotyledon expansion [228,229]; and GAMyb (gibberellin-induced myb-like transcription factor) that regulates the expression of α-amylase, mobilizing the starch during seed germination [214,230] (Figure 5).

The role of ROS in gibberellin signaling appears to be more related to seed germination, as it was demonstrated that hydrogen peroxide accelerates the germination and stimulates the early growth of seedlings [231]. In aleurone cells of cereal seeds, gibberellin decreases the activities of antioxidant enzymes SOD, CAT, and APX, contributing to the increase in hydrogen peroxide levels [232], which are involved in programmed cell death (PCD) during seed germination [233]. The ROS accumulation and PCD induced by GA in aleurone cells are suppressed by ABA [234].

6.3. Gibberellins and ROS on Drought Response

In stress conditions, the bioactive form of gibberellin is inactivated due to the positive regulation of GA2ox6 and GA2ox7, resulting in 2β-hydroxylation, leading to the activation of the DELLA repressor [235]. The reduction in gibberellin levels and accumulation of DELLA has been associated with increased drought tolerance. Reduced gibberellin levels were shown to trigger the activation of various stress-related genes [236] as well as the accumulation of osmolytes [237] and enzymes that scavenge ROS [238], all of which are linked to drought tolerance. Drought tolerance in tomatoes (Solanum lycopersicum), for example, can be achieved by decreasing the expression of gibberellin biosynthesis genes through the action of DREB protein [239], by the loss of the GID1 receptor gene [240], or by stable gain-of-function mutations in the RGA gene [241,242]. Similarly, transgenic tomatoes that overexpress arabidopsis GAMT1, responsible for methylating and inactivating gibberellin, exhibit reduced levels of gibberellin and increased drought tolerance [211]. Gibberellin signaling is an important regulator of cellular redox status, inhibiting the activity of various antioxidant enzymes [232]. Thus, the disruption of gibberellin during drought stress may play a key role in enhancing protection against oxidative stress, while ROS levels increase to fulfill their signaling functions.

7. Auxin

Auxin regulates cell division, elongation, tissue differentiation, embryogenesis, root formation, apical dominance, phyllotaxis, and tropic responses [243].

7.1. Auxin Metabolism

The indole-3-acetic acid (IAA) is the predominant naturally occurring auxin and is synthesized from the tryptophan precursor. Two steps compose the most important and well-characterized auxin biosynthesis pathway in plants, the TAA/YUC (tryptophan amino transferase/YUCCA) pathway. The amino acid tryptophan is first converted to indole-3-pyruvate (IPA) by the TAA (EC 2.6.1.99). Subsequently, the YUC-type flavin-containing monooxygenases (FMOs; EC 1.14.13.8) catalyze the production of IAA using IPA as a substrate [244], and this reaction seems to be the rate-limiting step in auxin biosynthesis [245]. IAA levels can be controlled mainly through IAA–amino acid conjugation by GH3 (Gretchen Hagen 3) or irreversible oxidation and degradation by DAO (dioxygenase for auxin oxidation) [246]. Auxin is synthesized at the shoot apex and transported by the mature phloem through a cell-to-cell transport mechanism named polar auxin transport [247]. Auxin flux in and out of the cell is mediated by AUX1/LAX (AUXIN RESISTANT1/LIKE AUX 1) permeases and PIN carrier proteins [248,249].

7.2. Auxin Signaling

Auxin-mediated signaling is dependent on the AUX/IAA (AUXIN/IAA) inhibitory proteins such as SLR1 (SOLITARY-ROOT 1) [250], AXR2 (AUXIN-RESISTANT 2) [251], and AXR3 (AUXIN-RESISTANT 3) [252]. AUX/IAA proteins contain an N-terminal EAR (ETHYLENE-RESPONSIVE ELEMENT BINDING FACTOR-ASSOCIATED AMPHIPHILIC REPRESSION) motif and a C-terminal PB1 (PHOX AND BEM 1) domain. In the absence of auxin, AUX/IAA binds to ARF (AUXIN RESPONSE FACTOR) transcription factors through its PB1 domain and to TPL/TPR (TOPLESS/TOPLESS-RELATED) corepressor proteins by its EAR motif, resulting in the repression of ARF [253] (Figure 6).

The nuclear receptor TIR1/AFB (TRANSPORT INHIBITOR RESPONSE 1/AUXIN SIGNALING F-BOX), which is associated with SCF-type E3-ubiquitin ligases, recognizes auxin, so in the presence of the phytohormone TIR1/AFB, it binds to the AUX/IAA repressor protein, leading to its ubiquitination and proteolysis. This results in the release of the transcriptional activity of ARF transcription factors, leading to expression of downstream auxin-responsive genes [253,254]. ARFs bind directly to the auxin-responsive promoters by the AuxRE (auxin-responsive element), regulating their transcriptional activation or repression (Figure 6).

In addition to the Aux/IAA/SCF^TIR1/AFB pathway, AUX response is also mediated by ABP1 (auxin binding protein 1). The quest for auxin receptors has spanned decades, with the identification of ABP1 long before TIR1/AFB. Initial research identified an abp1 mutant as the auxin receptor. Later works have led to uncertainty about ABP1’s role [255]. Recent re-analysis of abp1 mutants has demonstrated a division between proponents of ABP1 and TIR1 as primary auxin receptors [256,257,258,259]. Recent studies have confirmed ABP1 as a crucial auxin receptor, particularly for vascular formation and regeneration, confirmed by the observation that abp1 mutants show defects in auxin-induced vasculature [260]. ABP1 predominantly resides in the ER but is also secreted to the apoplast, where it binds auxin at the acidic pH. In the presence of auxin, the secreted ABP1 interacts with TMKs (TRANSMEMBRANE KINASES), forming the ABP1–TMK1 module that mediates rapid auxin-induced phosphorylation events [260,261], activating ion fluxes at the plasma membrane in response to auxin stimuli [262,263,264] and the MAPK cascade [265,266] (Figure 6). ABP1 also modulates the expression of AUX/IAA genes, indicating its broader role in influencing the Aux/IAA/SCF^TIR1/AFB pathway [267].

The two auxin receptors, ABP1 and TIR1, regulate multiple targets, with their actions tailored to the specific developmental context. ABP1 is critical for auxin regulation of the cell cycle, regulating the G1/S transition [268]. In fact, abnormal cell divisions occur within the suspensor and during the initial phases of embryo development in the arabidopsis mutant lacking ABP1 [269]. Auxin also stimulates cell elongation by initiating an acidification process within the cell wall, mediated by the activation of plasma membrane AHA2 (autoinhibited H⁺-ATPase 2), which exports protons to the apoplast (Figure 6). The low pH in the apoplast triggers a cascade of reactions that facilitate the “loosening” of the cell wall structure. This loosening will allow the irreversible expansion of the cell wall, driven by turgor pressure, ultimately leading to cell elongation [270]. The activation of AHA2 also induces hyperpolarization of the membrane potential, resulting in the opening of voltage-dependent inward K⁺ channels [271,272]. The influx of K⁺ contributes to water uptake and turgor, allowing cell expansion. Moreover, besides directly stimulating their activity, auxin also prompts the expression of both AHA2 channel and K⁺ channel genes [272,273].

AHA2 activity is inhibited by its dephosphorylation via PP2C-D family of protein phosphatases, which are inhibited by SAUR (small auxin upregulated RNA) [274]. SAUR19 is induced by auxin and promotes cell expansion through binding to PP2C-D and releasing AHA2 activity [275]. AHA2, as well as K⁺ inward channels, are also triggered by ABP1 during the initial perception of auxin, but the mechanism remains elusive [270] (Figure 6).

Because auxin regulates growth by cell elongation or division, auxin tends to act downstream in the regulation of many growth processes, such as cotyledon growth, hypocotyl elongation, root elongation, and lateral root formation [270,276]. Additionally, auxin biosynthesis, transport, and signaling are required by the ABA inhibitory effect in seed germination. This was demonstrated in the mutants yuc1 yuc6 [277], pin2 [278], and tir1 [279], which display resistance to the inhibitory effects of ABA on seed germination.

7.3. Auxin and ROS on Drought Response

Many works have highlighted the versatile involvement of auxins in enhancing plant tolerance to abiotic stresses [280]. Indeed, the overexpression of the auxin biosynthesis gene YUC6 is associated with enhanced drought tolerance and distinct phenotypic changes in arabidopsis [281], while the overexpression of OsGH3.2, involved in salicylic acid biosynthesis in rice, results in reduced levels of auxin and more sensitivity to drought [282,283]. In arabidopsis submitted to drought stress, auxin accumulation can also be triggered by downregulation of miRNA167. This miRNA targets IAR3 (IAA–ALANINE-RESISTANT 3) mRNA, which encodes an IAA–alanine hydrolase responsible for releasing active IAA from the IAA–alanine conjugate [284].

Auxin displays a crucial role in regulating genes involved in the response to water stress via ARF induction [285]. In arabidopsis, auxin positively regulates the expression of various stress-related genes, such as RAB18 (RESPONSIVE TO ABA 18), RD22 (RESPONSIVE TO DEHYDRATION 22), RD29A (RESPONSIVE TO DESICCATION 29A), RD29B (RESPONSIVE TO DESICCATION 29B), DREB2A, and DREB2B [286]. While ABA signaling induces stomatal closure, auxin regulates both stomatal movement and development, reducing stomatal density [287]. Arabidopsis mutants defective in auxin biosynthesis, transport, or signaling exhibit increased stomatal index, thereby establishing a negative role of auxin in stomatal development [288,289,290]. In mesophyll, auxin-induced ARF inhibits the expression of STOMAGEN peptide, which promotes stomata development [290,291]. During drought, plants utilize hydrotropism, an adaptive mechanism involving directed root growth towards water sources, to cope with this challenge. In this condition, the drought-induced auxin accumulation regulates the root system architecture, optimizing water uptake efficiency [292].

Auxin also contributes to the regulation of ROS levels, which are notably increased during drought. The RBOHs NADH oxidases’ expression and ROS levels are induced by RSL4 (ROOT HAIR DEFECTIVE SIX-LIKE 4), which is activated by ARF transcription factor and binds to at least one root hair cis-acting element (RHE) region in the RBOHC and RBOHJ promoters [293]. Indeed, ROS triggered by auxin are directly implicated in the loosening of the cell wall and contribute to the process of cell elongation [294]. The drought-induced ROS production significantly disrupts auxin homeostasis in plants [6], influencing polar auxin transport by modulating the expression PIN proteins and affecting auxin conjugation [295]. Similarly, ROS contributes to the oxidative degradation of auxin through peroxidase activity, resulting in changes in auxin gradients and disruption in hormone-mediated signaling [296]. Additionally, under water stress conditions, auxin has a positive regulatory effect on antioxidant enzymes SOD CAT, POD, and GR. This regulatory action intensifies ROS detoxification, thereby contributing to an effective defense mechanism against oxidative stress [286].

8. Cytokinin

Cytokinins are a group of phytohormones that regulate shoot and root growth, cell division and proliferation, chloroplast and vascular development, bud differentiation, anti-aging, biomass distribution [297,298,299], photosynthesis and nutrient metabolism [300], leaf senescence [301], and maintenance of meristem function [302,303]. In plants, the most abundant cytokinins are adenine-type species, including common derivatives such as isopentenyl adenine (iP), trans-zeatin (tZ), and cis-zeatin (cZ) [304]. In higher plants, zeatin is the most abundant cytokinin existing in both tZ and cZ forms. The tZ form is active in all plant species, whereas cZ is active only in specific plants, such as rice [305].

8.1. Cytokinin Metabolism

Cytokinins are synthesized via two primary biosynthetic pathways. The first one is de novo synthesis, where 5-phosphate adenosine (AMP, ATP, and ADP) is converted into iP/tZ-type cytokinins by adenosine phosphate isopentenyltransferase (IPT, EC 2.5.1.27) and CYP735A1/CYP735A2. The second biosynthetic pathway involves tRNA-specific adenylate isopentenyltransferase (tRNA-IPT), which indirectly leads to the formation of cZ-type cytokinins [304,306]. Cytokinins are synthesized in various cell types in both roots and shoots, and they undergo both local and long-distance transport [307,308,309,310]. Although the mechanisms of cytokinins transport are not fully understood, potential efflux and influx transporters have been identified, such as ABCG14, involved in cytokinin efflux across the plasma membrane [91,311].

Cytokinins degradation is primarily catalyzed by cytokinin oxidase/dehydrogenase (CKX, EC 1.5.99.12), resulting in the release of free adenine or adenine nucleosides, leading to the complete inactivation of the cytokinin [307]. Additionally, cytokinin inactivation can occur through conjugation to sugars via glucosyltransferases [304]. CKX and glucosyltransferases are both induced by cytokinins to deactivate the cytokinin signal itself. These negative-feedback pathways illustrate the tight regulation of cytokinin pathway output, reflecting its profound effects on plant development and the frequent targeting of cytokinin signaling by pathogens [91].

8.2. Cytokinin Signaling

Cytokinins signal transduction is analogous to the bacterial two-component regulatory system, which consists of histidine kinases (HKs; sensing proteins) and reactive regulatory proteins (receiving proteins) [312]. Cytokinins bind to the histidine kinases receptors AHK2 (ARABIDOPSIS HISTIDINE KINASE 2), AHK3, AHK4/CRE1 (CYTOKININ RESPONSE 1), CKI1 (CYTOKININ-INDEPENDENT 1), and AHK5/CKI2 to form a dimer [313]. The conserved His residues in the N-terminal sensor-kinase domain are autophosphorylated, and the phosphate group is then transferred to a conserved Asp in the C-terminal receiver domain [314,315,316,317,318]. In general loss-of-function mutations of single receptors displayed only subtle effects on most of the phenotypes studied. However, in some cases, individual receptors were shown to mediate specific cytokinin activities [319]. The histidine kinase receptors are primarily localized at the ER membrane, suggesting that the site of cytokinin binding is within the ER lumen [320,321,322]. A small portion of HK receptors are also found at the plasma membrane [323] although there is ongoing debate regarding the extent to which these receptors are involved in mediating the cytokinin response [311] (Figure 7).

During cytokinins signaling, the phosphate group on the conserved Asp residue of the receptor is subsequently transferred to HPt (histidine phosphate transfer) proteins in the cytoplasm. Cytokinin receptors interact in vivo with all studied HPt proteins and vice versa [324] (Figure 7). The arabidopsis genome contains five AUTHENTIC HISTIDINE-CONTAINING PHOSPHOTRANSMITTER genes: AHP1, AHP2, AHP3, AHP4, and AHP5. The AHP proteins feature a conserved cysteine residue susceptible to S-nitrosylation by nitric oxide, both in vitro and in vivo, that inhibits their ability to receive phosphate from histidine kinases and transfer phosphate to response regulators [325]. Additionally, plants possess pseudo-histidine phosphotransferase proteins (PHPs), which are lacking in the histidine phosphorylation site [326,327,328,329]. PHPs act as negative regulators of cytokinin signaling in arabidopsis, influencing protoxylem differentiation [330,331], leaf phyllotaxy [332], and cell patterning during lateral root initiation [333].

In the nucleus, phosphorylated HPT transfers phosphate groups to ARR (Arabidopsis response regulator), thereby regulating the expression of cytokinin-responsive genes [334]. In cytokinins signaling, two types of ARR play distinct roles. Type-B ARRs are phosphorylated on an Asp residue in their receiver domain by AHPs, crucial for initiating the transcriptional response to cytokinin [335,336]. Turnover rates within the type-B RR family vary and are regulated in part by the KISS ME DEADLY (KMD) E3–ubiquitin ligase complex [337,338]. Conversely, type-A ARRs function as negative-feedback regulators of cytokinin signaling. They are transcriptionally induced by cytokinin through direct activation by type-B ARRs [339,340,341,342]. Among the targets of the transcriptional control mediated by type-B RRs is the transcription factor WUSCHEL, which regulates shoot meristem activity, thereby establishing a direct link between cytokinin signaling and the control of shoot growth and development [343,344,345,346].

The CRF (CYTOKININ RESPONSE FACTOR) group also acts as a side-branch of the two-component signaling system, being activated by HPt proteins [347]. The CRF is a non-monolithic group of AP2/ERF transcription factors found in most angiosperms and appears to be a target of ARR-Bs [348]. CRFs are recognized as key transcription factors in responding to abiotic stresses across various plant species, contributing to the trade-off between growth and stress response [349,350]. Moreover, several members of the CRF family from different plant species are induced by oxidative stress, which is considered one of the most critical consequences of abiotic stress [351,352].

Cytokinins frequently synergize with other hormones, particularly auxin, to coordinate cell division and differentiation. CRFs are capable of binding to elements found in the promoters of some PIN genes, controlling the expression of auxin transporters [353] and IAA distribution during root development [354,355,356,357]. Additionally, CRF2 was identified as being a downstream target of ARF5/MONOPTEROS, a key auxin-regulated transcriptional factor [358], suggesting that CRF acts as an important bridge between cytokinin and auxin signaling. In the shoot, cytokinins promote cell proliferation in apical and axillary meristems. Increased cytokinins levels, resulting from reduced expression of CKX genes, have been linked to enhanced grain yield in indica rice varieties [359], and disruption of CKX genes in arabidopsis leads to increased seed yield [360]. Additionally, cytokinins regulate leaf phyllotaxy [332], gynoecium development [361,362,363,364], female gametophyte development [365,366], and vascular cambial development [367,368]. In contrast, cytokinins inhibit root growth by promoting cell differentiation in the root apical meristem and by suppressing both lateral root initiation and primary root elongation [369,370,371,372,373,374]. Indeed, cytokinins determine the size of arabidopsis root meristems by regulating the rate of cell differentiation in meristems [375]. Overexpression of type-B ARRs can promote arabidopsis root growth, indicating that cytokinins participate in root development [376]. In rice, the overexpression of OsIPT disrupts root development [377]. Similarly, overexpression of OsRR3 or OsRR5 improves root growth and induces lateral root initiation [378]. Conversely, overexpressing the Zea mays CKX gene in Nicotiana tabacum reduces cytokinin levels, inhibits stem development, and enhances root growth [371].

Cytokinins also collaborate with salicylic acid-regulated plant immunity by the interaction between ARR2 and the SA response factor TGA3, which binds to the PR1 promoter in arabidopsis during pathogen response [379]. Indeed, elevated cytokinin levels boost SA-mediated defense responses, providing greater resistance to infection by virulent oomycete pathogens [380] and arabidopsis transgenic plants overexpressing AtCRF5 enhance resistance to Pseudomonas syringae pv. tomato DC3000 (Pst DC3000) [381]. Additionally, DELLA proteins, acting as central nodes in transmitting environmental cues, directly engage with type-B ARRs to co-activate gene targets crucial for root growth and photomorphogenesis [382].

8.3. Cytokinin and ROS on Drought Response

Cytokinins can also enhance resistance to adverse environmental factors [300]. Different studies suggest that cytokinins signaling is directly involved in drought stress responses. Exogenous cytokinin application was shown to improve the drought resistance of Cucumis sativus [383]. Similarly, the expression of the cytokinin biosynthetic enzyme IPT in tobacco enhances drought tolerance by delaying leaf senescence [384]. The balance between triggering leaf senescence and sustaining photosynthesis is crucial for drought tolerance and for maintaining crop yields under stress [385,386,387].

In arabidopsis, the CRF family may affect cytokinins signaling in the equilibrium between active photosynthesis and senescence. AtCRF6 acts as a negative regulator of leaf senescence. Its overexpression preserves chlorophyll (Chl) and delays leaf yellowing [388]. Similarly, AtCRF9 promotes chlorophyll retention in dark-induced senescence assays, maintaining photosynthesis during senescence [389]. In the opposite direction, the overexpression of AtCRF1, AtCRF3, and AtCRF5 genes leads to early leaf senescence, and crf1,3,5,6 multiple knockout delays this process [390], indicating that these genes act as positive regulators of leaf senescence. The overexpression of AtCRF2 also accelerates senescence in rosette leaves [391], but it has also been shown to accelerate chloroplast division, a feature linked to enhanced photosynthetic activity [392,393].

Cytokinin is also demonstrated to play negative regulatory roles in drought stress response. In arabidopsis, cytokinin-deficient CKX-overexpressing plants show improved survival under drought stress [394]. Additionally, the receptors AHK2 and AHK3 [395]; the HPt proteins AHP2, AHP3, and AHP5 [396]; and the transcription factors ARR1, ARR10, and ARR12 [397] act as negative regulators of plant responses to drought. In tomato, cytokinins affect transpiration by regulating stomatal density and leaf size. Under drought stress, reduced levels of CKs lead to a decrease in the cell division rate, suppressing growth and reducing stomatal density. This, in turn, leads to lower transpiration rates, which enhances plant survival in drought conditions [398]. The cytokinin-mediated stomatal responses involve the apoplastic peroxidases PRX4, PRX33, PRX34, and PRX71 but not the NADPH oxidases RBOHD and RBOHF. While PRX2 acts as an ROS scavenger, PRX33, PRX34, and PRX71 contribute to ROS production and stomata closure. In guard cells, PRX2 expression is repressed by ARR2, while PRX33 is induced [160].

Stomata closure and ROS production induced by tZ are compromised in the histidine kinase ahk3 and response regulators arr2 mutants, which are also defective in FLG22 (flagellin 22)-mediated ROS production. On the other hand, IPT3 and ARR2 overexpression lines show higher levels of ROS in guard cells and constitutive closed stomata phenotypes [160]. Importantly, ost1-3 and aba1-3 mutants, which are defective in FLG22-triggered stomatal closure [399,400], show wild-type stomatal closure in response to tZ treatment.

9. Brassinosteroids

Steroid hormones have been implicated in several aspects of animal growth and development. In plants, the steroid-like hormone brassinosteroid (BR) plays important roles in plant growth and development, being involved in the control of several developmental and physiological processes, such as cell elongation, cell division, vascular differentiation, senescence, flowering time control, male fertility, pollen development, seed size, photomorphogenesis, and resistance to biotic and abiotic stresses [401]. Despite having been named due to its original identification in Brassica napus pollen [402], the most active brassinosteroid, brassinolide (BL), has been found widely distributed in virtually all plant species [403].

9.1. Brassinosteroids Metabolism

Many of the important advances in the study of brassinosteroids come from a series of different genetic screenings. An important step was the identification of brassinosteroids biosynthesis mutants such as de-etiolated 2 (det2) and constitutive photomorphogenesis (cpd). They were the first mutants identified in this biosynthesis pathway, and the genes encode, respectively, a 5α-reductase (EC 1.3.1.22) and a C23-steroid hydroxylase (EC 1.14.13.112) [404,405]. Both mutants have been identified in screening for mutants defective in hypocotyl elongation, and their phenotype can be reduced upon BL application. Brassinosteroids are produced in the cell, but several lines of evidence have shown that their perception occurs at the plasma membrane by a receptor complex that initiates the signaling [406]. Due to their polar nature, the bioactive brassinosteroids might be exported out the cells, but the mechanism by which this transportation occurs was previously unknown. Very recently, an arabidopsis ABC transporter protein (ABCB19) was identified as a functional brassinosteroid exporter [407].

9.2. Brassinosteroids Signaling

A deeper comprehension of the hormone’s perception and function comes from the identification of arabidopsis brassinosteroid signaling mutants. These mutants usually display dwarf phenotypes unable to respond to exogenous BL treatment. Brassinosteroids signaling is mediated by BR1 (BRASSINOSTEROID-INSENSITIVE 1) receptor, a membrane-bound receptor serine/threonine kinase protein with extracellular leucine-rich repeats [408,409,410]. Later, similar receptors were isolated and characterized in several other species, such as rice [411], tomato [412], pea (Pisum sativum) [413], barley (Hordeum vulgare) [414], currant tomato (Solanum pimpinellifolium) [415], petunia (Petunia hybrida) [416], wild tobacco (Nicotiana attenuate) [417], Brachypodium distachyon [418], maize (Zea mays) [419], and alfalfa (Medicago truncatula) [420]. The arabidopsis genome encodes three other BRI1-related genes. BRL1 and BRL3 but not BRL2 encode functional brassinosteroid receptors that bind BL with high affinity. These genes display different tissue-specific expressions. BRI1 is expressed ubiquitously in growing cells, while BRL1 and BRL3 expression are restricted to non-overlapping regions of the vascular tissue [421]. Similar to arabidopsis, BRI1-related genes have been also described in other plant species [411,414,419].

In the absence of brassinosteroid, the BRI1 receptor remains inactivated by several mechanisms (Figure 8). The cytoplasmatic C-terminal tail of BRI1 auto-inhibits its kinase activity [422], and the kinase domain is dephosphorylated by protein phosphatase 2A (PP2A), targeting the BRI1 pool for degradation through a negative-feedback loop [423]. PP2A is composed of a subset of cytoplasmatic β’ regulatory subunits (β′η, β′γ, β′ζ, and β′θ) [424] and is activated by methylation via SBI (SUPRESSOR OF BRI1), a leucine carboxylmethyltransferase (LCMT), reducing receptor abundance and BR signaling strength [423]. Another important negative regulation is performed by the inhibitory protein BKI1 (BRI1 kinase inhibitor 1), which binds to BRI1 and inhibits its activity [425].

BRI1 can bind brassinosteroids at the extracellular island domain [426], inducing its homodimerization and kinase activity [421,427,428,429], releasing the effect of the C-terminal tail [430], and leading to the trans-phosphorylation of the BKI1 inhibitor [425]. BKI1 dissociates from the membrane and interacts with members from the 14-3-3 phosphopeptide-binding proteins (Figure 8), allowing brassinosteroid signaling to be activated [425,431,432]. BRI1 then associates with another LRR receptor kinase, BAK1 (BRI1-associated kinase 1) [433,434] (Figure 8), also known as SERK3 (somatic embryogenesis receptor kinase 3). BAK1/SERK3 works as a brassinosteroid co-receptor, forming heterodimers with BRI1 [435,436] in the presence of the hormone [437]. Brassinosteroid-induced BRI1-BAK1 dimerization results in the positioning of their intracellular kinase domains in the right structure for competing with BKI1 and allows receptors’ transphosphorylation at multiple sites [427,438], forming a fully activated receptor complex that activates downstream events [427,438] (Figure 8). Two other members of the BAK1/SERK family, SERK1 [439] and SERK4/BAK7/BKK1 [440], have also been reported to be involved in BR signaling. Once fully activated, BRI1 phosphorylates CDG1 (CONSTITUTIVE DIFFERENTIAL GROWTH 1) and BSKs (BR SIGNALING KINASEs) proteins [438,439,440,441,442], both belonging to the receptor-like cytoplasmic kinase (RLCK) superfamily. These proteins are located at the plasma membrane by amino-terminal myristoylation [406]. Once phosphorylated by BRI1, CDG and BSK1 kinases phosphorylate the phosphatase BSU1 (BRI1 suppressor 1) [443,444]. BSU1 was identified in a bri1 suppressor screening [445]. BSU1 and its homologues, the BSU-likes (BSLs), belong to a small family of protein phosphatases known as PPKL (protein phosphatase with kelch-like domains) [445]. Once activated, they can dephosphorylate and inactivate GSK3-like kinase BIN2 (BRASSINOSTEROID-INSENSITIVE 2), a negative regulator of the brassinosteroid signaling [444,446] (Figure 8). Although BSU1 was the first gene to be functionally characterized and named in the family, it is specific to the Brassicaceae family and shows remarkable sequence variation, even among closely related species [447]. BSL genes, on the other hand, are highly conserved in all land plants [447].

BIN2 is a constitutively active kinase that acts as a central regulator of the brassinosteroid signaling cascade [443,448] (Figure 8). In arabidopsis, BIN2 and two close homologues, BIL1 (BRZ-INSENSITIVE-LONG 1) and BIL2, act redundantly in BR signaling since the triple loss-of-function mutant bin2-3 bil1 bil2 showed constitutive BR responses [448]. Once dephosphorylated by BSU and BSL, BIN2 becomes unstable, is degraded by a proteasome-mediated degradation [449], and releases the signal transduction cascade to its activated state (Figure 8). BIN2 activity is also regulated by interaction with the F-box E3 ligase KIB1 (KINK SUPPRESSED IN BZR1-1D), which, in the presence of brassinosteroids, will promote BIN2 ubiquitination and proteasomal degradation (Figure 8). KIB1 also blocks BIN2 access to its substrate, the BES1 (BRI1 EMS SUPPRESSOR 1)/BZR1 (BRASSINAZOLE-RESISTANT 1) transcriptional factors [450]. When the brassinosteroid levels are low or absent, the active BIN2 will hyperphosphorylate the two key transcriptional factors involved in the brassinosteroid response BES1 and BZR1 (Figure 8). Both genes have been isolated as gain-of-function dominant mutants and suppress weak bri1 mutant [451,452]. These mutants accumulate dephosphorylated BES1 and BZR1 proteins in the nucleus independently of brassinosteroids and display constitutive BR responses [451,452,453]. In their phosphorylated forms, BES1 and BZR1 cannot bind to DNA [454] and are retained at the cytoplasm by interaction with at least five of the twelve arabidopsis 14-3-3 proteins (14-3-3λ, 14-3-3κ.14-3-3ε, 14-3-3ϕ, and 14-4-3ω) [455] and the eight rice 14-3-3 proteins (GF14a-h; growth factor 14a–h) [456] (Figure 8).

In the opposite scenario, when the brassinosteroid levels are high, and BIN2 is inhibited, the BES1/BZR1 proteins are accumulated in their dephosphorylated forms (Figure 8). The dephosphorylation of BES1/BZR1 proteins is dependent on PP2A phosphatases’ activity [457]. Interestingly, the PP2A regulatory subunits involved in BES1/BZR1 dephosphorylation (PP2A β′α and PP2A β′β) are different from the ones involved in BRI1 dephosphorylation and are located at the nucleus [457]. The dephosphorylated forms of BES1/BZR1 will move to the nucleus, where they can bind to cis-acting elements in their target promoters, regulating gene expression [458,459,460,461] (Figure 8). In the arabidopsis genome, there are four other genes related to BES1 and BZR1, named BEH1–4 (BES1/BZR1 homologs 1–4), which have also been shown to operate in brassinosteroid signaling [458]. The promoters of the brassinosteroid biosynthesis genes display an enrichment in BRRE elements, where BES1 and BZR1 can bind as homodimers and repress its transcription, regulating both brassinosteroid biosynthesis and growth responses [459]. This operates as a negative-feedback loop to stop brassinosteroid signaling by the inhibition of the hormone production. Working as heterodimers with other transcriptional factors from the basic helix-loop-helix (bHLH) family, BES1 and BZR1 can bind to the E-box elements to activate brassinosteroid-induced gene expression [458]. The analysis of BL-induced gene promoters indicated that the E-box elements are enriched in many brassinosteroid-induced gene promoters [462].

BIM1 (BES1-interacting Myc-like 1) and its two homologues, BIM2 and BIM3, are members of the bHLH sub-family that are involved in brassinosteroid signaling. Although bim1, bim2, and bim3 single mutants do not display any visible phenotype, the bim1 bim2 bim3 triple mutants show a brassinosteroid mutant phenotype: shorter hypocotyls in light and dark growth conditions and higher sensitivity to brassinazole (a brassinosteroid biosynthesis inhibitor) in hypocotyl elongation assays in the dark [458]. On the other hand, BIM1 overexpression can partially rescue the bri1 mutant and can directly bind to the promoter of BR-induced genes [458]. Another transcription factor involved in the brassinosteroid response is known as BEE (brassinosteroid enhanced expression). In arabidopsis, the BEE genes are members of a subfamily of the bHLH, with 16 genes [463]. However, among the tested genes, only BEE1, BEE2, and BEE3 are responsive to brassinosteroid [464]. Similar to several other components of the brassinosteroid signaling that do not result in steroid-deficient mutant phenotypes (probably for gene duplication and functional redundancy), bee1, bee2, and bee3 single or double mutants do not display any visible phenotype. Only bee1 bee2 bee3 triple mutant has a reduced response to brassinosteroid and was shown to be shorter and display a light-grown phenotype similar to the weak brassinosteroid-response mutant but not the dwarf phenotype. This suggests that although BEE1, BEE2, and BEE2 are positive regulators, they are not absolutely required for brassinosteroid signaling [464].

Brassinosteroid signaling involves the critical step of signal attenuation. Excessive levels of BRI1 result in overly amplified brassinosteroid responses. To regulate this, plant U-box proteins PUB12 and PUB13 mediate BRI1 ubiquitination, triggering its internalization and subsequent degradation via the proteasome [465,466]. Signaling attenuation has been demonstrated to be an important step after brassinosteroid signaling. Since great amounts of BRI1 produce over-enhanced brassinosteroid responses, BRI1 internalization and degradation mediated by BRI1 ubiquitination by plant U-box proteins PUB12 and PUB13 have been shown to be an important step [465,466].

9.3. Brassinosteroids and ROS on Drought Response

Recent studies highlighted the importance of brassinosteroid signaling in drought adaptation [467]. In chickpea (Cicer arietinum), brassinosteroid treatment under water stress increased fresh and dry weight, the number of tillers, stem thickness, and root development [468]. In radish (Raphanus sativus) seedlings, it activated antioxidant enzymes, reducing drought stress effects [467]. In other species, such as sorghum (Sorghum bicolor), maize, and tomato, brassinosteroids enhanced chlorophyll accumulation, stomatal conductance, photosynthesis, and membrane stability, contributing to improved growth and yield [467].

In B. distachyon, BRI1-RNAi transgenic lines showed increased drought tolerance and increased expression of drought-induced genes [418]. The bri1 loss-of-function mutation led to drought resistance in arabidopsis [469,470]. Similar results have been observed in tomato, where a mutant in the SlBRI1 gene with weak brassinosteroid signaling (abs) exhibited drought tolerance. In comparison with control plants, abs mutants show a weak degree of wilting, higher water content in soil, and lower electrolyte leakage and MDA content. On the other hand, the overexpression of tomato SlBRI1 reduces stomatal aperture, antioxidant enzyme activities, and the expression of stress-related genes, negatively modulating drought tolerance [471]. In maize, under drought conditions, BRI1 RNAi-silenced transgenic lines display less wilting, higher survival rates, higher water content and water loss rate, higher stomatal closure, lower electrolyte leakage and MDA content, and a reduced photosynthetic decline rate in comparison with control plants. Opposite phenotypes have been observed in transgenic lines over-expressing ZmBRI1, indicating that ZmBR1 negatively modulates drought tolerance in maize [472]. These observations indicate that water stress regulates brassinosteroid signaling through BRI1 and that higher levels of the BRI1 receptor result in reduction in drought tolerance [470,471,472].

Overexpression of the BRL3 gene, another member of the brassinosteroid receptor family, enriched in the vascular system, confers tolerance to drought stress. Although loss-of-function mutations in the BRI1 receptor also lead to drought resistance, this occurs at the expense of plant growth. However, overexpression of the BRL3 receptor confers drought tolerance without penalizing overall growth. Systematic analyses have revealed that, under drought stress, increased BRL3 expression triggers the accumulation of osmoprotective metabolites, including proline and sugars [470]. Under control conditions, plants overexpressing the BRL3 receptor already exhibit a metabolic signature enriched in the accumulation of proline and sugars, which are classically correlated with stress tolerance. Under stress conditions, plants overexpressing BRL3 also accumulate high levels of proline, GABA, tyrosine, trehalose, myo-inositol, and raffinose in a manner correlated with stress exposure. All these metabolites have previously been associated with drought stress tolerance. High levels of the raffinose family oligosaccharides (RFOs) and myo-inositol, known to be involved in the protection of biological membranes and free radical scavenging, were also observed in the roots of plants overexpressing the BRL3 receptor. The results strongly suggest that BRL3 overexpression promotes the “priming” phenomenon, protecting plants from damage associated with the drought stress process [470].

OsGSK1, the rice orthologue of BIN2, was also shown to be involved in drought response. OsGSK1-knockout plants demonstrated more tolerance to several stresses, including drought [473]. In wheat (Triticum aestivum), a gain-of-function mutation in the TaGSK3 gene (TaGSK3-3D) confers drought tolerance. TaGSK3 encodes a SHAGGY-like kinase protein homologous to the arabidopsis BIN2. The gene was mapped and identified by the analysis of an Indian dwarf wheat cultivar (Sphaerococcum 1, S1) that is insensitive to brassinosteroid, displays upregulation of brassinosteroid biosynthetic genes, and enhances drouth tolerance [12]. TaGSK3-3D plants maintain higher relative water content through stomatal-regulated water loss and reduced ROS levels. Although TaGSK3-3D plants exhibited growth penalties under normal conditions, upon stress treatment, they displayed higher survival rates, plant height, and biomass compared to wild-type plants [474]. BIN2 and its homologues limit plant growth by inhibiting BES1/BZR1 activity; however, they enhance drought tolerance by phosphorylating SnRK2s, intensifying ABA signaling, and regulating other transcription factors involved in the activation of drought tolerance [475,476].

Under normal conditions, BES1 and the transcription factors WRKY46, WRKY54, and WRKY70 interact to promote the expression of genes related to growth and, at the same time, repress genes regulated by drought. On the other hand, drought stress conditions increase BES1 degradation, reducing its impact in plant growth and increasing drought response. In this condition, BIN2 phosphorylates WRKY54, allowing the expression of drought-inducible genes and reducing the growth stimulated by brassinosteroids [430]. BES1 also operates with the transcription factors RD26 (RESPONSIVE TO DESICCATION 26) [469] and TINY [477], coordinating plant growth and drought tolerance trade-off. RD26 and TINY are induced by drought and induce drought-responsive genes, increasing drought tolerance and inhibiting plant growth by inhibiting BES1. RD26 and TINY physically interact with BES1, inhibiting its transcriptional activity on brassinosteroid-regulated genes. On the other hand, BES1 represses the transcription of RD26, inhibiting drought responses [469,477]. BIN2 phosphorylates and stabilizes TYNY, performing a positive regulation in drought stress response [477].

In arabidopsis, a null mutant of the BEH3 gene (pca41, proline content alterative 41) shows a drought-insensitive phenotype, higher levels of proline accumulation, and reduced levels of reactive oxygen species. Under osmotic stress, the mutant displays an increase in APX, CAT, and POD activities. Overexpression of BEH3 results in an osmotic stress-sensitive phenotype that can be reverted by brassinolide application. BEH3 seems to operate together with the E3 ligase RZF1, and their expression mediates the water deficit response through brassinosteroid signaling and ubiquitination action [478]. Overexpression of the soybean BEH orthologue (GmBEH3L1) in arabidopsis also generates an osmotic-stress sensitive phenotype [479].

Interestingly, brassinosteroids signaling was also demonstrated as positively regulating drought stress. In wheat, the overexpression of TaBZR2 results in drought-tolerant plants, while downregulation of the gene by RNAi generates plants that exhibit elevated drought sensitivity. Under drought conditions, TaBZR2 RNAi plants display increased superoxide content, and it was demonstrated that TaBZR2 confers drought tolerance by activating TaGST1 expression [480]. Recently, a new gene involved in brassinosteroid signaling was identified through a genetic screening for a brassinazole-insensitive phenotype. Overexpression of BIL9 (BRZ-INSENSITIVE LONG HYPOCOTYL 9) leads to drought tolerance, while bil9-knockout plants do not present any visible phenotype under drought conditions. BIL9 overexpression induces the expression of ABA-induced and drought-responsive genes. BIL9 physically interacts with HOMEODOMAIN GLABROUS 11/ENHANCED DROUGHT TOLERANCE 1 (HDG11/EDT1), a transcription factor that promotes drought tolerance. BIL9/HDG11 interaction positively regulates BR-induced plant growth, promoting drought stress resistance [481].

Brassinosteroid signaling is also mediated by ROS. Brassinosteroid-induced hydrogen peroxide accumulation has been reported to be important for heat and oxidative stresses [482,483], stomatal movement [100,484], and salt tolerance [485]. Indeed, several components of the brassinosteroid signaling have been shown to be targets for ROS. The brassinosteroid co-receptor BAK1 was shown to be modified by glutaredoxin GRXC2, resulting in the inhibition of its kinase activity [486]. The kinase activity of BIN2 was also reported to be inhibited by nitric oxide [40]. Hydrogen peroxide leads to oxidative modifications in specific cysteine residues of BES1/BZR1, enhancing its transcriptional activity and promoting its interactions with other regulators of auxin and light signaling, such as ARF6 and PIF4. Oxidized BES1/BZR1 can be reduced by the thioredoxin TRXh5 [487]. Under hydrogen peroxide-deficient conditions, the number of genes regulated by BZR1 is greatly reduced, and the re-application of hydrogen peroxide partially restores the expression of the BZR1-regulated genes. Interestingly, BZR1 seems to repress ROS-related genes, contributing to feedback regulation and redox homeostasis [487]. Drought stress induces hydrogen peroxide production by brassinosteroid-activated NAPDPH oxidase, which may also activate BES1/BZR1 to regulate drought response [488].

Brassinosteroids signaling plays a pivotal role in balancing the trade-off between plant growth and drought tolerance. These hormones promote cell expansion and developmental processes under optimal conditions, enhancing overall growth. However, under drought stress, brassinosteroids trigger a complex adaptive response that prioritizes survival overgrowth. By modulating key signaling pathway, including those mediated by other phytohormones, brassinosteroids help plants adjust their metabolism, boost antioxidant defenses, and stabilize cellular structures, thereby minimizing the damage caused by water deficiency. As such, brassinosteroids are essential regulators that fine-tune the plant’s ability to conserve resources, optimizing both growth and stress resilience in challenging environments.

10. Strigolactones

Strigolactones are a small class of carotenoid-derived compounds that play a crucial role in suppressing shoot branching by inhibiting the outgrowth of axillary buds [489,490]. Initially, they were characterized as rhizosphere signals, helping root-parasitic plants detect and colonize their hosts [491]. However, strigolactones are now known to be involved in many other aspects of plant development [492]. They influence internode length [493,494], leaf morphology [495,496], leaf senescence [494,497], shoot gravitropism [498], stem thickness [499], as well as seed germination and early seedling development [500,501].

In the root system, strigolactones enhance the growth of primary roots, elongate root hairs, and promote the growth of rice crown roots [502,503,504]. Conversely, they inhibit adventitious root formation in eudicots [505,506,507]. Additionally, strigolactones play crucial roles in plant adaptive responses to environmental factors such as phosphate, nitrogen, light, drought, and high salinity [506,508,509,510,511,512].

10.1. Strigolactones Metabolism

In arabidopsis, MORE AXILLARY GROWTH genes MAX1, MAX3, and MAX4 encode enzymes involved in the strigolactone-biosynthetic pathway. MAX1 encodes a cytochrome P450 monooxygenase that is believed to be involved in a catalytic step downstream of MAX3 and MAX4 [509]. MAX3 and MAX4 encode carotenoid cleavage dioxygenase 7 (CCD7, EC 1.13.11.68) and CCD8 (EC 1.13.11.68), respectively. These enzymes catalyze sequential carotenoid cleavage reactions to produce an apocarotenone called carlactone, proposed as strigolactone precursor [513]. Indeed, an arabidopsis mutant in MAX3 and MAX4 exhibited a 70–75% reduction in strigolactones content [489]. After biosynthesis, a portion of strigolactones is exuded from the roots and enters the rhizosphere. In petunia, the PLEIOTROPIC DRUG RESISTANCE 1 (PDR1) protein played a crucial role in regulating the development of arbuscular mycorrhiza fungi and axillary branches [514]. PDR1 belongs to the ABC transporter family, which is also involved in the transport of other phytohormones, such as abscisic acid (ABA), auxin, and brassinosteroids [248,407,515,516]. Despite this, the mechanism underlying the transport of active strigolactones to the shoot remains unclear.

10.2. Strigolactones Signaling

The perception of strigolactones requires a hormone-dependent interaction between the receptor D14 and the F-box proteins D3 (in rice) or MAX2 (in arabidopsis), which are F-box leucine-rich repeat proteins structurally similar to the auxin receptor TIR1 and part of the SCF ubiquitination complex. During this process, strigolactones are hydrolyzed into a covalently linked intermediate molecule (CLIM), triggering a conformational change in D14 that facilitates its interaction with D3/MAX2 [517] (Figure 9). This mechanism is distinct from that of all known active phytohormones, which are produced by biosynthesis enzymes and then reversibly bound by their receptors to initiate signal transduction.

Strigolactone signaling depends on the inhibitory proteins SMAX1 (suppressor of MAX2 1), SMXL6 (SMAX-like 6), SMXL7 (SMAX-like 7), and SMXL8 (SMAX-like 8). In the absence of strigolactones, SMAX/SMXL proteins interact with transcriptional corepressors TPL and TPRs, potentially repressing the activities of target transcription factors such as BRC1 (BRANCHED1) [518,519,520] (Figure 9). The SMAX/SMXL proteins share high similarity with rice D53 and are also referred to as D53-like SMXLs [518,521]. In the presence of strigolactone, D3/MAX2 acts as the substrate-recruiting subunit of the SCF-type ubiquitin–E3 ligase complex D14-SCFD3 [509], which leads to the degradation of the inhibitory protein D53 through the 26S proteasome system, thereby allowing the expression of genes related to strigolactone response, such as BRC1, which encodes a TCP-type transcription factor that acts downstream of MAX2 in the regulation of shoot branching [509].

Strigolactones also interacts with other hormonal signaling. It has also been demonstrated that strigolactones induce the expression of CYTOKININ OXIDASISE DEHYDROGENASE 9 (CKX9), promoting cytokinin degradation [522,523]; inhibit auxin transport [524]; and regulate PIN auxin efflux carriers [525]. In apple, strigolactones regulate shoot regeneration through interaction with other phytohormones [526]. Strigolactones treatment induces members of the auxin biosynthesis gene family Yucca as well as Aux/IAA and ARF transcriptional factors, suggesting that strigolactones increase auxin levels, inhibiting shoot regeneration. The hormonal treatment also inhibits the ABA receptor PYL4 and the cytokinin oxidase GA2ox2 involved in the inhibition of active GA. Treatments with an inhibitor of strigolactones increase the expression of the brassinosteroid receptor BRI1 [526]. These results suggest that, besides the reduction in cytokinin and increased auxin signaling, abscisic acid, gibberellin, and brassinosteroids might also be involved in the strigolactone-mediated inhibition of shoot regeneration [526].

10.3. Strigolactones and ROS on Drought Response

Strigolactones positively regulate drought and high-salinity responses in arabidopsis. Both strigolactone-deficient and strigolactone-responsive mutants exhibit hypersensitivity to drought and salt stress. Exogenous strigolactone treatment rescues the drought-sensitive phenotype of strigolactone-deficient mutants but not of strigolactone-responsive mutants. This treatment also enhances drought tolerance in WT plants, confirming the role of strigolactones as positive regulators in stress responses. Additionally, the MAX3 and MAX4 genes, which are involved in strigolactone biosynthesis, are significantly induced by dehydration in leaves [527].

Strigolactones promote the expression of stress-related genes RD22, RD29a, and RD29b, improving tolerance to drought [528]. In addition, strigolactones are also involved in fine-tuning the plant’s response to oxidative stress by enhancing the expression and activity of antioxidant enzymes such as SOD, CAT, and POD [529,530] and promoting the production of non-enzymatic antioxidants, including reduced ascorbate, glutathione, and phenolic compounds, which scavenge ROS and protect against oxidative damage [531,532]. This helps maintain a balanced cellular redox state and shields plants from oxidative stress induced by environmental factors [529]. Thus, during the stress response, strigolactones play a critical role in the balance between ROS detoxification and production, ensuring the activation of ROS-dependent signaling and protecting against oxidative stress. However, as the field of strigolactones research is still evolving, further studies are needed to elucidate the precise mechanisms of strigolactones action and their potential applications in agricultural systems.

11. Phytohormones Crosstalk During Drought Response

Although ABA is recognized as the main phytohormone related to drought, the outcomes of response and tolerance to adverse conditions result from a complex interaction of multiple hormonal actions [533]. The stress response pathways exhibit intricate connections, often converging on common elements, which is referred to as “crosstalk”. The term crosstalk denotes situations where different signaling share one or more elements, resulting in shared outputs [534] (Figure 10). The plants’ physiological responses to stressful environments, involving changes in biological processes, arise from antagonistic or synergistic interactions between various phytohormones [535]. Consequently, the development of tolerance in plants against drought emerges as a complex phenomenon involving intricate interactions across various cellular, molecular, and metabolic dimensions [8].

The ABA regulation of stomatal closure under abiotic stress is directly related to ethylene signaling, which can act as either a positive or negative regulator [101]. This mechanism is dependent on ABA concentration. When ABA is not sufficient, drought stress leads to ethylene synthesis, which then induces the accumulation of ROS synergistically with ABA, inducing stomatal closure through the synthesis of NO and the activation of the SLACs channel [98,536]. In this condition, ethylene is also able to induce ABA biosynthesis through ERF transcription factors. In different species, the overexpression of ERFs results in increased ABA content, rapid stomal closure, and enhanced abiotic stress tolerance [101]. On the other hand, ABA inhibits ERF1 gene expression through a negative-feedback mechanism [537]. When ABA levels are high, ethylene starts to have an oppositive effect, inhibiting ABA-induced stomatal closure [98]: both exogenous ethylene and the ethylene-overproducing mutant eto1-1 show a reduced ABA-induced stomatal closure in drought-stressed arabidopsis [538]. It has been suggested that, under these conditions, ethylene might trigger the production of flavonoids, which could decrease NADPH oxidase activity. This would subsequently inhibit the synthesis of ROS induced by ABA, ultimately preventing ABA-induced stomatal closure [539,540]. Ethylene also regulates ROS metabolism by modulating antioxidant enzymes such as SOD, CAT, APX, and GR [541,542]. Similarly, the ethylene precursor ACC induces APX, CAT, SOD, and POX activities and reduces lipid peroxidation [543].

Despite the initial observation of water stress-induced ethylene synthesis [544], some studies have yielded varied and sometimes contradictory results, suggesting that drought stress either fails to increase or reduces ethylene production [545]. The ABA-induced transcription factor ABI4 mediates the transcriptional repression of ACS4 and ACS8, resulting in inhibition of ethylene biosynthesis during elevated ABA levels [98]. This is supported by the increased ethylene production in maize and tomato mutants deficient in ABA synthesis [546,547]. The combined ethylene–ABA stimulus results in half-open stomata, with diminished closure compared to the effect of the individual ABA or ethylene stimulus [548]. In this context, it was demonstrated that the inhibition of ethylene biosynthesis and perception can increase stomata closure, enhancing drought tolerance. In maize, the silencing of ACS decreases ethylene biosynthesis and improves drought tolerance [549]. Similarly, plants with reduced ethylene sensitivity, such as arabidopsis and maize overexpressing ARGOS genes [74] and arabidopsis mutants etr1-1 and ein2-1 [550], showed reduced water loss through transpiration and increased drought tolerance.

Despite this, reduced stomatal conductance and transpiration are not directly associated with drought tolerance, and the limitation of stomatal conductance is recognized as the primary factor contributing to the reduction in photosynthesis and oxidative stress during drought response [551,552,553]. Although reduced stomatal conductance leads to lower transpiration and water loss, it also restricts the supply of CO₂ for RubisCO (EC 4.1.1.39) carboxylase activity, thereby limiting photosynthesis and diverting ribulose bisphosphate (RuBP) to photorespiration, which is the primary source of ROS in photosynthetic tissues [551,552,553]. Therefore, the regulation of ABA-induced stomatal closure, along with the activation of the antioxidant system through the ethylene signaling, plays a central role in ensuring an appropriate response to drought (Figure 10).

Under moderate drought conditions, ABA can positively regulate root system architecture, increasing primary root length and root hair density through its induction of auxin biosynthesis and transport in the primary roots [554,555,556,557]. Indeed, the knockout of the ABA catabolic gene ABA8ox in rice [556] or the exogenous ABA application in arabidopsis [555] increases the auxin flux towards the root, resulting in increased root length and drought tolerance. Low concentrations of ABA also facilitate primary root growth by mitigating PP2C-D-mediated inhibition of the apoplastic efflux of H⁺ through the AHA2 channel [558]. For effective hydrotropism, the ABA-induced kinase SnRK2.2 is required at the cortical cells in the root elongation zone, where it promotes cellular elongation, allowing differential growth [559]. On the other hand, at elevated concentrations, ABA inhibits root growth by disrupting auxin signaling in roots, repressing genes encoding transporters responsible for auxin translocation [560]. Taken together, these observations show that both auxin and ABA play crucial roles in shaping the root system architecture in response to water availability. Mild soil drying and low ABA concentrations stimulate root growth by inducing auxin signaling. However, as drought conditions intensify, and ABA and ROS levels increase, auxin signaling and root growth are suppressed. While this inhibition does not enhance water absorption, it serves as an adaptive strategy during severe drought, allowing plants to allocate resources toward preserving photosynthetic tissues, thereby maximizing their chances of survival [561].

ABA and brassinosteroids are known to act antagonistically, balancing growth and stress response. BIN2 kinase seems to be the central hub of this interaction since its activity is repressed by dephosphorylation mediated by PP2Cs, a part of the central core of ABA signaling [476,562]. Under drought conditions, the increase in ABA leads to the repression of the PP2Cs ABI1 and ABI2. This allows BIN2 to be an active kinase, attenuating growth responses due to the phosphorylation of BES1/BZR1 transcriptional factors, which reduces the transcription of the brassinosteroid-induced genes related to growth [476]. On the other hand, BIN2 also phosphorylates SnRK2s, intensifying ABA signaling [475,563]. ABA also negatively regulates the expression of the transcriptional factors BEE1, BEE2, and BEE3 [464].

Pioneer analyses of auxin and brassinosteroid co-regulation have demonstrated that the signaling of both hormones converges at the promoters of shared target genes [462,564]. BES1 and BZR1 also integrate brassinosteroid signaling with other phytohormone signaling, co-regulating plant growth and stress tolerance [565,566,567,568]. Promoter regions from several transcriptional factor genes involved in light, auxin, and gibberellin signaling are targets of BZR1 [460]. BES1 and BZR1 also interact with PIF and DELLA transcriptional factors, coregulating several genes and modulating cell elongation and photomorphogenesis [456,569,570,571].

The presence of jasmonic acid in the roots has been suggested as essential during water stress to increase ABA levels [27,572]. Despite this, jasmonic acid inhibits root growth and reduces meristem activity [573]. This inhibition appears to be mediated by the interaction between jasmonic acid and auxin signaling [574]. Upon jasmonic acid induction, MYC2 binds to the promoters of the auxin-responsive gene PLT (PLETHORA), responsible for stem cell niche maintenance and cell division, leading to suppression of its expression and thereby inhibiting root meristem activity [574]. On the other hand, jasmonic acid inhibition of root growth is negatively regulated by brassinosteroids [575], which act antagonistically to jasmonic acid during plant defense [576]. The antagonistic regulation of metabolic genes is the main feature of gibberellin and ABA interactions [574]. ABA-deficient mutants exhibit elevated gibberellin levels, highlighting ABA’s role in suppressing gibberellin metabolic genes (GA3ox1/2 and GA3ox1/2/3) during seed germination. Conversely, gibberellin-deficient mutants show increased ABA levels by upregulating ABA biosynthetic genes (ABA1, NCED6, and NCED9) while downregulating the ABA catabolic gene (CYP707A2) [577,578]. Under water-limited conditions, gibberellin can inhibit ABA biosynthesis and stomatal closure [579]. Gibberellin is also recognized to antagonize ABA regulation of different developmental stages, including seed dormancy, seed germination, root growth, leaf development, flowering time, and responses to environmental cues such as light, temperature, and other abiotic stresses [580].

The gibberellin signaling appears to impair stress responses. The DELLA protein family, which inhibits gibberellin signaling in its absence, also promotes an ABA response by promoting the expression of the ABA transporter gene ABA-IMPORTING TRANSPORTER 1.1 (AIT1.1) in guard cells [540] and activating ABI transcription factors [581]. Transgenic tomato plants overexpressing the constitutively active DELLA protein proceraΔ17 (proΔ17) showed decreased stomatal aperture and plant transpiration; however, these effects are suppressed in the ABA-deficient sitiens (sit) mutant, suggesting DELLA’s role is ABA-dependent [241]. Similarly, DELLA can bind and inhibit JAZ, the central negative regulator of jasmonic acid signaling [582]. Gibberellin-mediated degradation of DELLA contributes to improve JAZ inhibition of jasmonate-responsive gene expression [583,584] and jasmonate-mediated plant immune responses [585].

Despite antagonizing the stress-induced ABA and jasmonate responses, gibberellin acts synergically to auxin, brassinosteroids, and ethylene, exerting a pivotal role in the modulation of intricated molecular and cellular processes that allow plants to respond to environmental conditions [239]. Gibberellin can regulate the auxin transport by inducing the expression of the PIN-FORMED auxin transporters PIN1, PIN2, and PIN3. This mechanism has an important role in gravitropism and xylogenesis [294,586]. Indeed, arabidopsis gibberellin biosynthesis- and signaling-deficient mutants show reduced activity of PIN, and the wild-type phenotype is restored upon exogenous gibberellin application [587]. Additionally, DELLA protein RGA interacts with and inhibits the auxin-responsible factors ARF6, ARF7, and ARF8. Gibberellin enhances the transcription of auxin response by promoting DELLA degradation [568]. Similar mechanisms appear to allow gibberellin to positively regulate the transcription of brassinosteroids and ethylene-responsive genes. DELLA also regulates BES1 and BZR1 1transcription factors, which control the responses and are activated by gibberellin via DELLA degradation [588], indicating an important intersection between gibberellin and brassinosteroids signaling. In ethylene signaling, the degradation of DELLA mediated by gibberellin is accompanied by the inhibition of the stress-responsive genes AP2/ERF [239]. Additionally, gibberellin induces ethylene biosynthesis by inducing ACS5 and ACS8 [589].

Stress-induced ABA accumulation downregulates cytokinin biosynthesis through the MYB2 transcription factor, which relieves the repression of cytokinin signaling and activates ABA- and stress-inducible genes [590]. Indeed, ABA and cytokinins regulate bud outgrowth antagonistically [591]. Under drought conditions, tZ transport is impaired [592]. Key kinases in the ABA signaling, such as SnRK2.2, SnRK2.3, and SnRK2.6, directly phosphorylate several Ser residues of ARR5, a type-A ARR and inhibitor of cytokinin signaling. This phosphorylation stabilizes the ARR5 protein, enhancing drought tolerance by suppressing cytokinin signaling and by positively regulating ABA signaling in an SnRK2-dependent manner [593]. Despite acting antagonistically, during biotic stress response, cytokinins induce ROS production and stomata closure. This mechanism is not affected in the ost1-3 mutant, indicating that it occurs in an ABA-independent manner [160]. In arabidopsis, both ABA and drought downregulate the expression of ARR1, ARR10, and ARR12 [397]. Additionally, ABA-activated ABI4 binds the promoters of ARR6, ARR7, and ARR15, further impairing their expression [594,595,596]. Thus, the ABA-mediated inhibition of cytokinin signaling can reshape the plant body by downregulating shoot growth while accelerating root growth. This allows the plant to reduce water loss and increase water uptake from deeper soil layers [590].

Conversely, elevated endogenous cytokinins suppress ABA signaling, thereby influencing a trade-off between growth and defense mechanisms. This reduced sensitivity to ABA under high cytokinins levels is thought to be mediated through type-B ARRs such as ARR1, ARR11, and ARR12, which interact directly with SnRK2s, inhibiting the kinase activity of SnRK2.6 [593]. Indeed, AHK2, AHK3, and AHK4 defective mutants as well as type-B ARRs ARR1, ARR10, and ARR16 exhibit higher sensitivity to ABA and display increased resistance to drought [397,597,598]. On the other hand, the type-A ARRs ARR4, ARR5, and ARR6 downregulate ABI5 expression, contributing to ABA response [595].

Strigolactones act as positive regulators of stress signaling networks and various ABA signaling by regulating the expression of many stress- and ABA-responsive genes involved in plant development and abiotic stress responses. Impaired strigolactone signal transduction also leads to the downregulation of CKX genes, which are necessary for cytokinin degradation [527]. These findings suggest that coordinated crosstalk between strigolactones, ABA, and cytokinins signaling networks regulates plant adaptation to adverse environmental conditions. Consistent with the drought-sensitive phenotype, max mutants, which show disrupted strigolactone biosynthesis, displayed increased leaf stomatal density compared to wild-type plants and exhibited slower ABA-induced stomatal closure. Relative to the wild type, max mutants showed a higher rate of leaf water loss during dehydration and reduced ABA responsiveness during germination and post germination, underscoring the role of strigolactones as positive regulators of ABA signaling [527].

12. Concluding Remarks

In summary, plant hormones play a crucial and multifaceted role in mediating plant responses to drought stress. Unlike animal hormones, which often have a single, well-defined function, plant hormones operate across a spectrum of functions, with their effects depending on the interplay between different signaling. This complexity allows plants to finely tune their responses to the timing and intensity of drought stress.

ABA is recognized as the primary regulator of drought responses, driving stomatal closure through ROS production and optimizing root tissue patterning to minimize water loss and enhance uptake. Ethylene complements ABA at low concentrations but opposes its effects when ABA levels are high. Salicylic acid also supports ABA by inducing ROS-mediated stomatal closure when ABA signaling is insufficient; jasmonates improve ABA accumulation, aiding stomatal closure and root adaptations; and strigolactones enhance drought tolerance by amplifying ABA responses, regulating stress-related genes, and strengthening antioxidant defenses. In contrast, gibberellins and cytokinins counteract ABA by suppressing stomatal closure, ROS production, and growth under stress. Auxins and brassinosteroids act mainly in the balance between growth and stress adaptation, optimizing root architecture and enhancing antioxidant defenses (Figure 10).

These nine classes of plant hormones interact synergistically, coordinating a comprehensive response to stress while balancing growth and survival. These hormones do not function in isolation but rather through intricate networks where crosstalk between pathways is vital. Notably, ROS often mediate these signaling, acting as key molecular signals that integrate stress responses (Figure 10). These mechanisms are essential for plant stress responses and occur differently depending on the intensity of the stress. In moderate stress conditions, such as mild drought, the crosstalk between various signaling promotes coordinated responses that enable the plant to maintain growth and adapt to the environment, such as auxin activation to stimulate root growth (Figure 10). However, under severe stress, like intense drought, these crosstalk mechanisms shift, prioritizing the preservation of photosynthetic tissues and minimizing cellular damage. The trade-off between growth and stress response reflects the differential allocation of energy resources, allowing the plant to prioritize survival under stress without irreversibly compromising development (Figure 10). In this context, the interaction between different phytohormones and ROS signaling can inhibit processes such as root growth, favoring adaptive strategies that ensure the plant’s survival by redirecting resources to essential vital functions.

From an evolutionary perspective, the ability of plants to modulate their responses through such diverse and interconnected hormonal signals has been crucial for their adaptation to fluctuating environments. As we face increasing climatic extremes and the pressing need to expand agricultural production to meet growing food demands, understanding these hormonal pathways offers the potential to identify new biotechnological targets. This knowledge is essential for developing crops with enhanced resilience to drought, ultimately contributing to global food security and sustainable agricultural practices.

Author Contributions

Conceptualization, D.J.-M. and G.S.-M.; formal analysis, D.J.-M., Y.d.S.-V. and G.S.-M.; resources, D.J.-M. and G.S.-M.; writing—original draft preparation, D.J.-M., Y.d.S.-V. and G.S.-M.; writing—review and editing, D.J.-M. and G.S.-M.; visualization, D.J.-M. and G.S.-M.; supervision, D.J.-M. and G.S.-M.; project administration, D.J.-M. and G.S.-M.; funding acquisition, D.J.-M. and G.S.-M. All authors have read and agreed to the published version of the manuscript.

Acknowledgments

We are grateful to Ana Lucia de Moraes Giannini for critically reading the manuscript. We would like to dedicate this review to Joanne Chory (1955–2024), for her inspiring work on plant hormonal signalling and her pioneer work in how plants respond to the environmental changes, a legacy that inspired and will continuing to inspire generations.

Conflicts of Interest

The authors declare no conflicts of interest.

Footnotes

Disclaimer/Publisher’s Note: The statements, opinions, and data contained in all publications are solely those of the individual author(s) and contributor(s) and not of MDPI and/or the editor(s). MDPI and/or the editor(s) disclaim responsibility for any injury to people or property resulting from any ideas, methods, instructions, or products referred to in the content.

Footnotes

Disclaimer/Publisher’s Note: The statements, opinions and data contained in all publications are solely those of the individual author(s) and contributor(s) and not of MDPI and/or the editor(s). MDPI and/or the editor(s) disclaim responsibility for any injury to people or property resulting from any ideas, methods, instructions or products referred to in the content.

Figures

References

1. Hamann, E.; Denney, D.; Day, S.; Lombardi, E.; Jameel, M.I.; MacTavish, R.; Anderson, J.T. Plant eco-evolutionary responses to climate change: Emerging directions. Plant Sci.; 2021; 304, 110737. [DOI: https://dx.doi.org/10.1016/j.plantsci.2020.110737]

2. Ahuja, I.; de Vos, R.C.H.; Bones, A.M.; Hall, R.D. Plant molecular stress responses face climate change. Trends Plant Sci.; 2010; 15, pp. 664-674. [DOI: https://dx.doi.org/10.1016/j.tplants.2010.08.002] [PubMed: https://www.ncbi.nlm.nih.gov/pubmed/20846898]

3. Tigchelaar, M.; Battisti, D.S.; Naylor, R.L.; Ray, D.K. Future warming increases probability of globally synchronized maize production shocks. Proc. Natl. Acad. Sci. USA; 2018; 115, pp. 6644-6649. [DOI: https://dx.doi.org/10.1073/pnas.1718031115] [PubMed: https://www.ncbi.nlm.nih.gov/pubmed/29891651]

4. Farooq, M.A.; Niazi, A.K.; Akhtar, J.; Saifullah,; Farooq, M.; Souri, Z.; Karimi, N.; Rengel, Z. Acquiring control: The evolution of ROS-Induced oxidative stress and redox signaling pathways in plant stress responses. Plant Physiol. Biochem.; 2019; 141, pp. 353-369. [DOI: https://dx.doi.org/10.1016/j.plaphy.2019.04.039] [PubMed: https://www.ncbi.nlm.nih.gov/pubmed/31207496]

5. Kumar, A.; Memo, M.; Mastinu, A. Plant behaviour: An evolutionary response to the environment?. Plant Biol.; 2020; 22, pp. 961-970. [DOI: https://dx.doi.org/10.1111/plb.13149]

6. Xia, X.J.; Zhou, Y.H.; Shi, K.; Zhou, J.; Foyer, C.H.; Yu, J.Q. Interplay between reactive oxygen species and hormones in the control of plant development and stress tolerance. J. Exp. Bot.; 2015; 66, pp. 2839-2856. [DOI: https://dx.doi.org/10.1093/jxb/erv089]

7. Salvi, P.; Manna, M.; Kaur, H.; Thakur, T.; Gandass, N.; Bhatt, D.; Muthamilarasan, M. Phytohormone signaling and crosstalk in regulating drought stress response in plants. Plant Cell Rep.; 2021; 40, pp. 1305-1329. [DOI: https://dx.doi.org/10.1007/s00299-021-02683-8]

8. Weyers, J.D.B.; Paterson, N.W. Plant hormones and the control of physiological processes. New Phytol.; 2001; 152, pp. 375-407. [DOI: https://dx.doi.org/10.1046/j.0028-646X.2001.00281.x] [PubMed: https://www.ncbi.nlm.nih.gov/pubmed/33862994]

9. Suzuki, N.; Rivero, R.M.; Shulaev, V.; Blumwald, E.; Mittler, R. Abiotic and biotic stress combinations. New Phytol.; 2014; 203, pp. 32-43. [DOI: https://dx.doi.org/10.1111/nph.12797]

10. Waadt, R.; Seller, C.A.; Hsu, P.K.; Takahashi, Y.; Munemasa, S.; Schroeder, J.I. Plant hormone regulation of abiotic stress responses. Nat. Rev. Mol. Cell Biol.; 2022; 23, pp. 680-694. [DOI: https://dx.doi.org/10.1038/s41580-022-00479-6]

11. Zheng, Y.; Wang, X.; Cui, X.; Wang, K.; Wang, Y.; He, Y. Phytohormones regulate the abiotic stress: An overview of physiological, biochemical, and molecular responses in horticultural crops. Front. Plant Sci.; 2023; 13, 1095363. [DOI: https://dx.doi.org/10.3389/fpls.2022.1095363] [PubMed: https://www.ncbi.nlm.nih.gov/pubmed/36684767]

12. Gupta, A.; Hua, L.; Lin, G.; Molnár, I.; Doležel, J.; Liu, S.; Li, W. Multiple origins of Indian dwarf wheat by mutations targeting the TREE domain of a GSK3-like kinase for drought tolerance, phosphate uptake, and grain quality. Theor. Appl. Genet.; 2021; 134, pp. 633-645. [DOI: https://dx.doi.org/10.1007/s00122-020-03719-5] [PubMed: https://www.ncbi.nlm.nih.gov/pubmed/33164159]

13. Zhu, J.-K. Abiotic stress signaling and responses in plants. Cell; 2016; 167, pp. 313-324. [DOI: https://dx.doi.org/10.1016/j.cell.2016.08.029]

14. Bandurska, H. Drought Stress Responses: Coping Strategy and Resistance. Plants; 2022; 11, 922. [DOI: https://dx.doi.org/10.3390/plants11070922]

15. Iqbal, S.; Wang, X.; Mubeen, I.; Kamran, M.; Kanwal, I.; Díaz, G.A.; Abbas, A.; Parveen, A.; Atiq, M.N.; Alshaya, H. et al. Phytohormones Trigger Drought Tolerance in Crop Plants: Outlook and Future Perspectives. Front. Plant Sci.; 2022; 12, 799318. [DOI: https://dx.doi.org/10.3389/fpls.2021.799318] [PubMed: https://www.ncbi.nlm.nih.gov/pubmed/35095971]

16. Sharkey, T.D.; Raschke, K. Effects of phaseic Acid and dihydrophaseic Acid on stomata and the photosynthetic apparatus. Plant Physiol.; 1980; 65, pp. 291-297. [DOI: https://dx.doi.org/10.1104/pp.65.2.291] [PubMed: https://www.ncbi.nlm.nih.gov/pubmed/16661176]

17. Osakabe, Y.; Yamaguchi-Shinozaki, K.; Shinozaki, K.; Tran, L.P. ABA control of plant macroelement membrane transport systems in response to water deficit and high salinity. New Phytol.; 2014; 202, pp. 35-49. [DOI: https://dx.doi.org/10.1111/nph.12613] [PubMed: https://www.ncbi.nlm.nih.gov/pubmed/24283512]

18. Corratgé-Faillie, C.; Lacombe, B. Substrate (un)specificity of Arabidopsis NRT1/PTR FAMILY (NPF) proteins. J. Exp. Bot.; 2017; 68, pp. 3107-3113. [DOI: https://dx.doi.org/10.1093/jxb/erw499]

19. Parveen, A.; Ahmar, S.; Kamran, M.; Malik, Z.; Ali, A.; Riaz, M.; Abbasi, G.H.; Khan, M.; Sohail, A.B.; Rizwan, M. et al. Abscisic acid signaling reduced transpiration flow, regulated Na+ ion homeostasis and antioxidant enzyme activities to induce salinity tolerance in wheat (Triticum aestivum L.) seedlings. Environ. Technol. Innov.; 2021; 24, 101808. [DOI: https://dx.doi.org/10.1016/j.eti.2021.101808]

20. Bhaskara, G.B.; Nguyen, T.T.; Verslues, P.E. Unique drought resistance functions of the highly ABA-induced clade a protein phosphatase 2Cs. Plant Physiol.; 2012; 160, pp. 379-395. [DOI: https://dx.doi.org/10.1104/pp.112.202408] [PubMed: https://www.ncbi.nlm.nih.gov/pubmed/22829320]

21. Hasan, M.M.; Liu, X.D.; Waseem, M.; Guang-Qian, Y.; Alabdallah, N.M.; Jahan, M.S.; Fang, X.W. ABA activated SnRK2 kinases: An emerging role in plant growth and physiology. Plant Signal. Behav.; 2022; 17, 2071024. [DOI: https://dx.doi.org/10.1080/15592324.2022.2071024]

22. Aleman, F.; Yazaki, J.; Lee, M.; Takahashi, Y.; Kim, A.Y.; Li, Z.; Kinoshita, T.; Ecker, J.R.; Schroeder, J.I. An ABA-increased interaction of the PYL6 ABA receptor with MYC2 Transcription Factor: A putative link of ABA and JA signaling. Sci. Rep.; 2016; 6, 28941. [DOI: https://dx.doi.org/10.1038/srep28941]

23. Park, S.Y.; Fung, P.; Nishimura, N.; Jensen, D.R.; Fujii, H.; Zhao, Y.; Lumba, S.; Santiago, J.; Rodrigues, A.; Chow, T.F. et al. Abscisic acid inhibits type 2C protein phosphatases via the PYR/PYL family of START proteins. Science; 2009; 324, pp. 1068-1071. [DOI: https://dx.doi.org/10.1126/science.1173041]

24. Feng, C.Z.; Chen, Y.; Wang, C.; Kong, Y.H.; Wu, W.H.; Chen, Y.F. Arabidopsis RAV1 transcription factor, phosphorylated by SnRK2 kinases, regulates the expression of ABI3, ABI4, and ABI5 during seed germination and early seedling development. Plant J.; 2014; 80, pp. 654-668. [DOI: https://dx.doi.org/10.1111/tpj.12670] [PubMed: https://www.ncbi.nlm.nih.gov/pubmed/25231920]

25. Fujita, Y.; Fujita, M.; Shinozaki, K.; Yamaguchi-Shinozaki, K. ABA-mediated transcriptional regulation in response to osmotic stress in plants. J. Plant Res.; 2011; 124, pp. 509-525. [DOI: https://dx.doi.org/10.1007/s10265-011-0412-3] [PubMed: https://www.ncbi.nlm.nih.gov/pubmed/21416314]

26. Furihata, T.; Maruyama, K.; Fujita, Y.; Umezawa, T.; Yoshida, R.; Shinozaki, K.; Yamaguchi-Shinozaki, K. Abscisic acid-dependent multisite phosphorylation regulates the activity of a transcription activator AREB1. Proc. Natl. Acad. Sci. USA; 2006; 103, pp. 1988-1993. [DOI: https://dx.doi.org/10.1073/pnas.0505667103] [PubMed: https://www.ncbi.nlm.nih.gov/pubmed/16446457]

27. de Ollas, C.; Dodd, I.C. Physiological impacts of ABA–JA interactions under water-limitation. Plant Mol. Biol.; 2016; 91, pp. 641-650. [DOI: https://dx.doi.org/10.1007/s11103-016-0503-6] [PubMed: https://www.ncbi.nlm.nih.gov/pubmed/27299601]

28. Goda, H.; Sasaki, E.; Akiyama, K.; Maruyama-Nakashita, A.; Nakabayashi, K.; Li, W.; Ogawa, M.; Yamauchi, Y.; Preston, J.; Aoki, K. et al. The AtGenExpress hormone and chemical treatment data set: Experimental design, data evaluation, model data analysis and data access. Plant J.; 2008; 55, pp. 526-542. [DOI: https://dx.doi.org/10.1111/j.1365-313X.2008.03510.x] [PubMed: https://www.ncbi.nlm.nih.gov/pubmed/18419781]

29. Finkelstein, R. Abscisic Acid Synthesis and Response. The Arabidopsis Book; U.S. National Library of Medicine: Bethesda, MA, USA, 2013; Volume 11, e0166. [DOI: https://dx.doi.org/10.1199/tab.0166]

30. Kwak, J.M.; Mori, I.C.; Pei, Z.M.; Leonhardt, N.; Torres, M.A.; Dangl, J.L.; Bloom, R.E.; Bodde, S.; Jones, J.D.G.; Schroeder, J.I. NADPH oxidase AtrbohD and AtrbohF genes function in ROS-dependent ABA signaling in Arabidopsis. EMBO J.; 2003; 22, pp. 2623-2633. [DOI: https://dx.doi.org/10.1093/emboj/cdg277] [PubMed: https://www.ncbi.nlm.nih.gov/pubmed/12773379]

31. Shen, J.; Zhang, J.; Zhou, M.; Zhou, H.; Cui, B.; Gotor, C.; Romero, L.C.; Fu, L.; Yang, J.; Foyer, C.H. et al. Persulfidation-based modification of cysteine desulfhydrase and the NADPH oxidase RBOHD controls guard cell abscisic acid signaling. Plant Cell; 2020; 32, pp. 1000-1017. [DOI: https://dx.doi.org/10.1105/tpc.19.00826] [PubMed: https://www.ncbi.nlm.nih.gov/pubmed/32024687]

32. Yoshida, T.; Fernie, A.R.; Shinozaki, K.; Takahashi, F. Long-distance stress and developmental signals associated with abscisic acid signaling in environmental responses. Plant J.; 2021; 105, pp. 477-488. [DOI: https://dx.doi.org/10.1111/tpj.15101]

33. Mittler, R.; Blumwald, E. The roles of ROS and ABA in systemic acquired acclimation. Plant Cell; 2015; 27, pp. 64-70. [DOI: https://dx.doi.org/10.1105/tpc.114.133090] [PubMed: https://www.ncbi.nlm.nih.gov/pubmed/25604442]

34. Hua, D.; Wang, C.; He, J.; Liao, H.; Duan, Y.; Zhu, Z.; Guo, Y.; Chen, Z.; Gong, Z. A plasma membrane receptor kinase, GHR1, mediates abscisic acid- and hydrogen peroxide-regulated stomatal movement in Arabidopsis. Plant Cell; 2012; 24, pp. 2546-2561. [DOI: https://dx.doi.org/10.1105/tpc.112.100107] [PubMed: https://www.ncbi.nlm.nih.gov/pubmed/22730405]

35. Roelfsema, M.R.G.; Hedrich, R.; Geiger, D. Anion channels: Master switches of stress responses. Trends Plant Sci.; 2012; 17, pp. 221-229. [DOI: https://dx.doi.org/10.1016/j.tplants.2012.01.009] [PubMed: https://www.ncbi.nlm.nih.gov/pubmed/22381565]

36. Kuromori, T.; Seo, M.; Shinozaki, K. ABA Transport and Plant Water Stress Responses. Trends Plant Sci.; 2018; 23, pp. 513-522. [DOI: https://dx.doi.org/10.1016/j.tplants.2018.04.001]

37. Wang, P.; Song, C.P. Guard-cell signalling for hydrogen peroxide and abscisic acid. New Phytol.; 2008; 178, pp. 703-718. [DOI: https://dx.doi.org/10.1111/j.1469-8137.2008.02431.x] [PubMed: https://www.ncbi.nlm.nih.gov/pubmed/18373649]

38. Zhao, C.; Cai, S.; Wang, Y.; Chen, Z.H. Loss of nitrate reductases NIA1 and NIA2 impairs stomatal closure by altering genes of core ABA signaling components in Arabidopsis. Plant Signal. Behav.; 2016; 11, e1183088. [DOI: https://dx.doi.org/10.1080/15592324.2016.1183088]

39. Castillo, M.C.; Lozano-Juste, J.; González-Guzmán, M.; Rodriguez, L.; Rodriguez, P.L.; León, J. Inactivation of PYR/PYL/RCAR ABA receptors by tyrosine nitration may enable rapid inhibition of ABA signaling by nitric oxide in plants. Sci. Signal.; 2015; 8, ra89. [DOI: https://dx.doi.org/10.1126/scisignal.aaa7981]

40. Wang, P.; Du, Y.; Hou, Y.J.; Zhao, Y.; Hsu, C.C.; Yuan, F.; Zhu, X.; Tao, W.A.; Song, C.P.; Zhu, J.K. Nitric oxide negatively regulates abscisic acid signaling in guard cells by S-nitrosylation of OST1. Proc. Natl. Acad. Sci. USA; 2015; 112, pp. 613-618. [DOI: https://dx.doi.org/10.1073/pnas.1423481112]

41. Liu, Y.; Shi, L.; Ye, N.; Liu, R.; Jia, W.; Zhang, J. Nitric oxide-induced rapid decrease of abscisic acid concentration is required in breaking seed dormancy in Arabidopsis. New Phytol.; 2009; 183, pp. 1030-1042. [DOI: https://dx.doi.org/10.1111/j.1469-8137.2009.02899.x] [PubMed: https://www.ncbi.nlm.nih.gov/pubmed/19522839]

42. Lozano-Juste, J.; León, J. Enhanced abscisic acid-mediated responses in nia1 nia2 noa1-2 triple mutant impaired in NIA/NR- and AtNOA1-dependent nitric oxide biosynthesis in Arabidopsis. Plant Physiol.; 2010; 152, pp. 891-903. [DOI: https://dx.doi.org/10.1104/pp.109.148023]

43. He, H.; Van Breusegem, F.; Mhamdi, A. Redox-dependent control of nuclear transcription in plants. J. Exp. Bot.; 2018; 69, pp. 3359-3372. [DOI: https://dx.doi.org/10.1093/jxb/ery130]

44. Matsuo, M.; Johnson, J.M.; Hieno, A.; Tokizawa, M.; Nomoto, M.; Tada, Y.; Godfrey, R.; Obokata, J.; Sherameti, I.; Yamamoto, Y.Y. et al. High REDOX RESPONSIVE TRANSCRIPTION FACTOR1 Levels Result in Accumulation of Reactive Oxygen Species in Arabidopsis thaliana Shoots and Roots. Mol. Plant; 2015; 8, pp. 1253-1273. [DOI: https://dx.doi.org/10.1016/j.molp.2015.03.011] [PubMed: https://www.ncbi.nlm.nih.gov/pubmed/25882345]

45. Scintu, D.; Scacchi, E.; Cazzaniga, F.; Vinciarelli, F.; De Vivo, M.; Shtin, M.; Svolacchia, N.; Bertolotti, G.; Unterholzener, S.J.; Del Bianco, M. et al. microRNA165 and 166 modulate response of the Arabidopsis root apical meristem to salt stress. Commun. Biol.; 2023; 6, 834. [DOI: https://dx.doi.org/10.1038/s42003-023-05201-6]

46. Ramachandran, P.; Augstein, F.; Mazumdar, S.; Nguyen TVan Minina, E.A.; Melnyk, C.W.; Carlsbecker, A. Abscisic acid signaling activates distinct VND transcription factors to promote xylem differentiation in Arabidopsis. Curr. Biol.; 2021; 31, pp. 3153-3161.e5. [DOI: https://dx.doi.org/10.1016/j.cub.2021.04.057] [PubMed: https://www.ncbi.nlm.nih.gov/pubmed/34043949]

47. Doubt, S.L. The response of plants to illuminating gas. Bot. Gaz.; 1917; 63, pp. 209-224. [DOI: https://dx.doi.org/10.1086/332006]

48. Nath, P.; Trivedi, P.K.; Sane, V.A.; Sane, A.P. Role of ethylene in fruit ripening. Ethylene Action in Plants; Khan, N. Springer: Berlin/Heidelberg, Germany, 2006; pp. 151-184.

49. Trusov, Y.; Botella, J.R. Silencing of the ACC synthase gene ACACS2 causes delayed flowering in pineapple [Ananas comosus (L.) Merr.]. J. Exp. Bot.; 2006; 57, pp. 3953-3960. [DOI: https://dx.doi.org/10.1093/jxb/erl167]

50. Wilmowicz, E.; Kesy, J.; Kopcewicz, J. Ethylene and ABA interactions in the regulation of flower induction in Pharbitis nil. J. Plant Physiol.; 2008; 165, pp. 1917-1928. [DOI: https://dx.doi.org/10.1016/j.jplph.2008.04.009] [PubMed: https://www.ncbi.nlm.nih.gov/pubmed/18565620]

51. Wuriyanghan, H.; Zhang, B.; Cao, W.H.; Ma, B.; Lei, G.; Liu, Y.F.; Wei, W.; Wu, H.J.; Chen, L.J.; Chen, H.W. et al. The ethylene receptor ETR2 delays floral transition and affects starch accumulation in rice. Plant Cell; 2009; 21, pp. 1473-1494. [DOI: https://dx.doi.org/10.1105/tpc.108.065391]

52. Etchells, J.P.; Provost, C.M.; Turner, S.R. Plant vascular cell division is maintained by an interaction between PXY and ethylene signalling. PLoS Genet.; 2012; 8, e1002997. [DOI: https://dx.doi.org/10.1371/journal.pgen.1002997] [PubMed: https://www.ncbi.nlm.nih.gov/pubmed/23166504]

53. Corbineau, F.; Xia, Q.; Bailly, C.; El-Maarouf-Bouteau, H. Ethylene, a key factor in the regulation of seed dormancy. Front. Plant Sci.; 2014; 5, 539. [DOI: https://dx.doi.org/10.3389/fpls.2014.00539] [PubMed: https://www.ncbi.nlm.nih.gov/pubmed/25346747]

54. Dubois, M.; Van den Broeck, L.; Inzé, D. The Pivotal Role of Ethylene in Plant Growth. Trends Plant Sci.; 2018; 23, pp. 311-323. [DOI: https://dx.doi.org/10.1016/j.tplants.2018.01.003]

55. Feng, G.; Sanderson, B.J.; Keefover-Ring, K.; Liu, J.; Ma, T.; Yin, T.; Smart, L.B.; DiFazio, S.P.; Olson, M.S. Pathways to sex determination in plants: How many roads lead to Rome?. Curr. Opin. Plant Biol.; 2020; 54, pp. 61-68. [DOI: https://dx.doi.org/10.1016/j.pbi.2020.01.004]

56. Clarke, S.M.; Cristescu, S.M.; Miersch, O.; Harren, F.J.M.; Wasternack, C.; Mur, L.A.J. Jasmonates act with salicylic acid to confer basal thermotolerance in Arabidopsis thaliana. New Phytol.; 2009; 182, pp. 175-187. [DOI: https://dx.doi.org/10.1111/j.1469-8137.2008.02735.x] [PubMed: https://www.ncbi.nlm.nih.gov/pubmed/19140948]

57. Thomashow, M.F. Molecular basis of plant cold acclimation: In sights gained from studying the CBF cold response pathway. Plant Physiol.; 2010; 154, pp. 571-577. [DOI: https://dx.doi.org/10.1104/pp.110.161794] [PubMed: https://www.ncbi.nlm.nih.gov/pubmed/20921187]

58. Du, H.; Wu, N.; Cui, F.; You, L.; Li, X.H.; Xiong, L.Z. A homolog of ETHYLENE OVERPRODUCER, OsETOL1, differentially modulates drought and submergence tolerance in rice. Plant J.; 2014; 78, pp. 834-849. [DOI: https://dx.doi.org/10.1111/tpj.12508]

59. Guan, R.; Su, J.B.; Meng, X.Z.; Li, S.; Liu, Y.D.; Xu, J.; Zhang, S.Q. Multilayered regulation of ethylene induction plays a positive role in Arabidopsis resistance against Pseudomonas syringae. Plant Physiol.; 2015; 169, pp. 299-312. [DOI: https://dx.doi.org/10.1104/pp.15.00659] [PubMed: https://www.ncbi.nlm.nih.gov/pubmed/26265775]

60. Tao, J.J.; Chen, H.W.; Ma, B.; Zhang, W.K.; Chen, S.Y.; Zhang, J.S. The role of ethylene in plants under salinity stress. Front. Plant Sci.; 2015; 6, 1059. [DOI: https://dx.doi.org/10.3389/fpls.2015.01059] [PubMed: https://www.ncbi.nlm.nih.gov/pubmed/26640476]

61. Perata, P. Ethylene signaling controls fast oxygen sensing in plants. Trends Plant Sci.; 2020; 25, pp. 3-6. [DOI: https://dx.doi.org/10.1016/j.tplants.2019.10.010]

62. Miyazaki, J.H.; Yang, S.F. Metabolism of 5-methylthioribose to methionine. Plant Physiol.; 1987; 84, pp. 277-281. [DOI: https://dx.doi.org/10.1104/pp.84.2.277] [PubMed: https://www.ncbi.nlm.nih.gov/pubmed/16665430]

63. Roje, S. S-adenosyl-L-methionine: Beyond the universal methyl group donor. Phytochemistry; 2006; 67, pp. 1686-1698. [DOI: https://dx.doi.org/10.1016/j.phytochem.2006.04.019] [PubMed: https://www.ncbi.nlm.nih.gov/pubmed/16766004]

64. Argueso, C.T.; Hansen, M.; Kieber, J.J. Regulation of ethylene biosynthesis. J. Plant Growth Regul.; 2007; 26, pp. 92-105. [DOI: https://dx.doi.org/10.1007/s00344-007-0013-5]

65. Hegg, E.L.; Que-Jr, L. The 2-his-1-carboxylate facial triad—An emerging structural motif in mononuclear non-heme iron(II) enzymes. Eur. J. Biochem.; 1997; 250, pp. 625-629. [DOI: https://dx.doi.org/10.1111/j.1432-1033.1997.t01-1-00625.x] [PubMed: https://www.ncbi.nlm.nih.gov/pubmed/9461283]

66. Rodrigues, M.A.; Bianchetti, R.E.; Freschi, L. Shedding light on ethylene metabolism in higher plants. Front. Plant Sci.; 2014; 5, 665. [DOI: https://dx.doi.org/10.3389/fpls.2014.00665] [PubMed: https://www.ncbi.nlm.nih.gov/pubmed/25520728]

67. Chang, C.; Kwok, S.F.; Bleecker, A.B.; Meyerowitz, E.M. Arabidopsis ethylene-response gene ETR1: Similarity of product to two-component regulators. Science; 1993; 262, pp. 539-544. [DOI: https://dx.doi.org/10.1126/science.8211181]

68. Hua, J.; Meyerowitz, E.M. Ethylene responses are negatively regulated by a receptor gene family in Arabidopsis thaliana. Cell; 1998; 94, pp. 261-271. [DOI: https://dx.doi.org/10.1016/S0092-8674(00)81425-7] [PubMed: https://www.ncbi.nlm.nih.gov/pubmed/9695954]

69. Qiao, H.; Chang, K.N.; Yazaki, J.; Ecker, J.R. Interplay between ethylene, ETP1/ETP2 F-box proteins, and degradation of EIN2 triggers ethylene responses in Arabidopsis. Genes Dev.; 2009; 23, pp. 512-521. [DOI: https://dx.doi.org/10.1101/gad.1765709] [PubMed: https://www.ncbi.nlm.nih.gov/pubmed/19196655]

70. Resnick, J.S.; Wen, C.K.; Shockey, J.A.; Chang, C. REVERSION-TOETHYLENE SENSITIVITY1, a conserved gene that regulates ethylene receptor function in Arabidopsis. Proc. Natl. Acad. Sci. USA; 2006; 103, pp. 7917-7922. [DOI: https://dx.doi.org/10.1073/pnas.0602239103]

71. Resnick, J.S.; Rivarola, M.; Chang, C. Involvement of RTE1 in conformational changes promoting ETR1 ethylene receptor signaling in Arabidopsis. Plant J.; 2008; 56, pp. 423-431. [DOI: https://dx.doi.org/10.1111/j.1365-313X.2008.03615.x]

72. Dong, C.H.; Jang, M.; Scharein, B.; Malach, A.; Rivarola, M.; Liesch, J.; Groth, G.; Hwang, I.; Chang, C. Molecular association of the Arabidopsis ETR1 ethylene receptor and a regulator of ethylene signaling, RTE1. J. Biol. Chem.; 2010; 285, pp. 40706-40713. [DOI: https://dx.doi.org/10.1074/jbc.M110.146605] [PubMed: https://www.ncbi.nlm.nih.gov/pubmed/20952388]

73. Rai, M.I.; Wang, X.; Thibault, D.M.; Kim, H.J.; Bombyk, M.M.; Binder, B.M.; Shakeel, S.N.; Schaller, G.E. The ARGOS gene family functions in a negative feedback loop to desensitize plants to ethylene. BMC Plant Biol.; 2015; 15, 157. [DOI: https://dx.doi.org/10.1186/s12870-015-0554-x] [PubMed: https://www.ncbi.nlm.nih.gov/pubmed/26105742]

74. Shi, J.; Habben, J.E.; Archibald, R.L.; Drummond, B.J.; Chamberlin, M.A.; Williams, R.W.; Lafitte, H.R.; Weers, B.P. Overexpression of ARGOS Genes Modifies Plant Sensitivity to Ethylene, Leading to Improved Drought Tolerance in Both Arabidopsis and Maize. Plant Physiol.; 2015; 169, pp. 266-282. [DOI: https://dx.doi.org/10.1104/pp.15.00780] [PubMed: https://www.ncbi.nlm.nih.gov/pubmed/26220950]

75. Hirayama, T.; Kieber, J.J.; Hirayama, N.; Kogan, M.; Guzman, P.; Nourizadeh, S.; Alonso, J.M.; Dailey, W.P.; Dancis, A.; Ecker, J.R. RESPONSIVE-TO ANTAGONIST1, a Menkes/Wilson disease-related copper transporter, is required for ethylene signaling in Arabidopsis. Cell; 1999; 97, pp. 383-393. [DOI: https://dx.doi.org/10.1016/S0092-8674(00)80747-3] [PubMed: https://www.ncbi.nlm.nih.gov/pubmed/10319818]

76. Binder, B.M.; Rodríguez, F.I.; Bleecker, A.B. The copper transporter RAN1 is essential for biogenesis of ethylene receptors in Arabidopsis. J. Biol. Chem.; 2010; 285, pp. 37263-37270. [DOI: https://dx.doi.org/10.1074/jbc.M110.170027]

77. Binder, B.M.; Walker, J.M.; Gagne, J.M.; Emborg, T.J.; Hemmann, G.; Bleecker, A.B.; Vierstra, R.D. The Arabidopsis EIN3 binding F-Box proteins EBF1 and EBF2 have distinct but overlapping roles in ethylene signaling. Plant Cell; 2007; 19, pp. 509-523. [DOI: https://dx.doi.org/10.1105/tpc.106.048140] [PubMed: https://www.ncbi.nlm.nih.gov/pubmed/17307926]

78. Müller, M.; Munné-Bosch, S. Ethylene Response Factors: A Key Regulatory Hub in Hormone and Stress Signaling. Plant Physiol.; 2015; 169, pp. 32-41. [DOI: https://dx.doi.org/10.1104/pp.15.00677]

79. Solano, R.; Stepanova, A.; Chao, Q.; Ecker, J.R. Nuclear events in ethylene signaling: A transcriptional cascade mediated by ethylene-insensitive 3 and ethylene-response-factor 1. Genes Dev.; 1998; 12, pp. 3703-3714. [DOI: https://dx.doi.org/10.1101/gad.12.23.3703]

80. Ju, C.; Yoon, G.M.; Shemansky, J.M.; Lin, D.Y.; Ying, Z.I.; Chang, J.; Garrett, W.M.; Kessenbrock, M.; Growth, G.; Tucker, M.L. et al. CTR1 phosphorylates the central regulator EIN2 to control ethylene hormone signalling from the ER membrane to the nucleus in Arabidopsis. Proc. Natl. Acad. Sci. USA; 2012; 109, pp. 19486-19491. [DOI: https://dx.doi.org/10.1073/pnas.1214848109] [PubMed: https://www.ncbi.nlm.nih.gov/pubmed/23132950]

81. Azhar, B.J.; Zulfiqar, A.; Shakeel, S.N.; Schaller, G.E. Amplification and adaptation in the ethylene signaling pathway. Small Methods; 2020; 4, 1900452. [DOI: https://dx.doi.org/10.1002/smtd.201900452]

82. Binder, B.M. Ethylene signaling in plants. J. Biol. Chem.; 2020; 295, pp. 7710-7725. [DOI: https://dx.doi.org/10.1074/jbc.REV120.010854]

83. Yoo, S.-D.; Cho, Y.-H.; Tena, G.; Xiong, Y.; Sheen, J. Dual control of nuclear EIN3 by bifurcate MAPK cascades in C₂H₄ signalling. Nature; 2008; 451, pp. 789-795. [DOI: https://dx.doi.org/10.1038/nature06543] [PubMed: https://www.ncbi.nlm.nih.gov/pubmed/18273012]

84. Hahn, A.; Harter, K. Mitogen-activated protein kinase cascades and ethylene: Signaling, biosynthesis, or both?. Plant Physiol.; 2009; 149, pp. 1207-1210. [DOI: https://dx.doi.org/10.1104/pp.108.132241] [PubMed: https://www.ncbi.nlm.nih.gov/pubmed/19109412]

85. Stepanova, A.N.; Alonso, J.M. Ethylene signaling and response: Where different regulatory modules meet. Curr. Opin. Plant Biol.; 2009; 12, pp. 548-555. [DOI: https://dx.doi.org/10.1016/j.pbi.2009.07.009]

86. Lin, Z.; Ho, C.W.; Grierson, D. AtTRP1 encodes a novel TPR protein that interacts with the ethylene receptor ERS1 and modulates development in Arabidopsis. J. Exp. Bot.; 2009; 60, pp. 3697-3714. [DOI: https://dx.doi.org/10.1093/jxb/erp209]

87. Scharein, B.; Voet-van-Vormizeele, J.; Harter, K.; Groth, G. Ethylene signaling: Identification of a putative ETR1-AHP1 phosphorelay complex by fluorescence spectroscopy. Anal. Biochem.; 2008; 377, pp. 72-76. [DOI: https://dx.doi.org/10.1016/j.ab.2008.03.015] [PubMed: https://www.ncbi.nlm.nih.gov/pubmed/18384742]

88. Urao, T.; Miyata, S.; Yamaguchi-Shinozaki, K.; Shinozaki, K. Possible His to Asp phosphorelay signaling in an Arabidopsis two-component system. FEBS Lett.; 2000; 478, pp. 227-232. [DOI: https://dx.doi.org/10.1016/S0014-5793(00)01860-3] [PubMed: https://www.ncbi.nlm.nih.gov/pubmed/10930573]

89. Zdarska, M.; Cuyacot, A.R.; Tarr, P.T.; Yamoune, A.; Szmitkowska, A.; Hrdinová, V.; Gelová, Z.; Meyerowitz, E.M.; Hejátko, J. ETR1 integrates response to ethylene and cytokinins into a single multistep phosphorelay pathway to control root growth. Mol. Plant; 2019; 12, pp. 1338-1352. [DOI: https://dx.doi.org/10.1016/j.molp.2019.05.012]

90. Scharein, B.; Groth, G. Phosphorylation alters the interaction of the Arabidopsis phosphotransfer protein AHP1 with its sensor kinase ETR1. PLoS ONE; 2011; 6, e24173. [DOI: https://dx.doi.org/10.1371/journal.pone.0024173]

91. Kieber, J.J.; Schaller, G.E. Cytokinin signaling in plant development. Development; 2018; 145, dev149344. [DOI: https://dx.doi.org/10.1242/dev.149344] [PubMed: https://www.ncbi.nlm.nih.gov/pubmed/29487105]

92. Street, I.H.; Aman, S.; Zubo, Y.; Ramzan, A.; Wang, X.; Shakeel, S.N.; Kieber, J.J.; Schaller, G.E. Ethylene inhibits cell proliferation of the Arabidopsis root meristem. Plant Physiol.; 2015; 169, pp. 338-350. [DOI: https://dx.doi.org/10.1104/pp.15.00415]

93. Fatma, M.; Asgher, M.; Iqbal, N.; Rasheed, F.; Sehar, Z.; Sofo, A.; Khan, N.A. Ethylene Signaling under Stressful Environments: Analyzing Collaborative Knowledge. Plants; 2022; 11, 2211. [DOI: https://dx.doi.org/10.3390/plants11172211]

94. Sharma, A.; Kumar, V.; Sidhu, G.P.S.; Kumar, R.; Kohli, S.K.; Yadav, P.; Kapoor, D.; Bali, A.S.; Shahzad, B.; Khanna, K. et al. Abiotic Stress Management in Plants: Role of Ethylene. Molecular Plant Abiotic Stress; Roychoudhury, A.; Tripathi, D. John Wiley & Sons: Hoboken, NJ, USA, 2019; pp. 185-208. [DOI: https://dx.doi.org/10.1002/9781119463665.ch10]

95. Oh, S.A.; Park, J.H.; Lee, G.I.; Paek, K.H.; Park, S.K.; Nam, H.G. Identification of three genetic loci controlling leaf senescence in Arabidopsis thaliana. Plant J.; 1997; 12, pp. 527-535. [DOI: https://dx.doi.org/10.1111/j.0960-7412.1997.00527.x]

96. Zhang, T.Y.; Li, Z.Q.; Zhao, Y.D.; Shen, W.J.; Chen, M.S.; Gao, H.Q.; Ge, X.M.; Wang, H.Q.; Li, X.; He, J.M. Ethylene-induced stomatal closure is mediated via MKK1/3-PK3/6 cascade to EIN2 and EIN3. J. Integr. Plant Biol.; 2021; 63, pp. 1324-1340. [DOI: https://dx.doi.org/10.1111/jipb.13083] [PubMed: https://www.ncbi.nlm.nih.gov/pubmed/33605510]

97. Ge, X.M.; Cai, H.L.; Lei, X.; Zhou, X.; Yue, M.; He, J.M. Heterotrimeric G protein mediates ethylene-induced stomatal closure via hydrogen peroxide synthesis in Arabidopsis. Plant J.; 2015; 82, pp. 138-150. [DOI: https://dx.doi.org/10.1111/tpj.12799] [PubMed: https://www.ncbi.nlm.nih.gov/pubmed/25704455]

98. Hasan, M.M.; Liu, X.D.; Yao, G.Q.; Liu, J.; Fang, X.W. Ethylene-mediated stomatal responses to dehydration and rehydration in seed plants. J. Exp. Bot.; 2024; 75, pp. 6719-6732. [DOI: https://dx.doi.org/10.1093/jxb/erae060]

99. Liu, J.; Hou, L.; Liu, G.; Liu, X.; Wang, X. Hydrogen sulfide induced by nitric oxide mediates ethylene-induced stomatal closure of Arabidopsis thaliana. Chin. Sci. Bull.; 2011; 56, pp. 3547-3553. [DOI: https://dx.doi.org/10.1007/s11434-011-4819-y]

100. Shi, C.; Qi, C.; Ren, H.; Huang, A.; Hei, S.; She, X. Ethylene mediates brassinosteroid-induced stomatal closure via Gα protein-activated hydrogen peroxide and nitric oxide production in Arabidopsis. Plant J.; 2015; 82, pp. 280-301. [DOI: https://dx.doi.org/10.1111/tpj.12815] [PubMed: https://www.ncbi.nlm.nih.gov/pubmed/25754244]

101. Müller, M. Foes or Friends: ABA and Ethylene Interaction under Abiotic Stress. Plants; 2021; 10, 448. [DOI: https://dx.doi.org/10.3390/plants10030448]

102. Desikan, R.; Hancock, J.T.; Bright, J.; Harrison, J.; Weir, I.; Hooley, R.; Neill, S.J. A role for ETR1 in hydrogen peroxide signaling in stomatal guard cells. Plant Physiol.; 2005; 137, pp. 831-834. [DOI: https://dx.doi.org/10.1104/pp.104.056994] [PubMed: https://www.ncbi.nlm.nih.gov/pubmed/15761208]

103. Vlot, A.C.; Dempsey, D.A.; Klessig, D.F. Salicylic Acid, a multifaceted hormone to combat disease. Annu. Rev. Phytopathol.; 2009; 47, pp. 177-206. [DOI: https://dx.doi.org/10.1146/annurev.phyto.050908.135202] [PubMed: https://www.ncbi.nlm.nih.gov/pubmed/19400653]

104. Klessig, D.F.; Choi, H.W.; Dempsey, D.A. Systemic acquired resistance and salicylic acid: Past, present, and future. Mol. Plant Microbe Interact.; 2018; 31, pp. 871-888. [DOI: https://dx.doi.org/10.1094/MPMI-03-18-0067-CR]

105. An, C.; Mou, Z. Salicylic acid and its function in plant immunity. J. Integr. Plant Biol.; 2011; 53, pp. 412-428. [DOI: https://dx.doi.org/10.1111/j.1744-7909.2011.01043.x] [PubMed: https://www.ncbi.nlm.nih.gov/pubmed/21535470]

106. Raskin, I.; Ehmann, A.; Melander, W.R.; Meeuse, B.J.D. Salicylic acid: A natural inducer of heat production in Arum lilies. Science; 1987; 237, pp. 1601-1602. [DOI: https://dx.doi.org/10.1126/science.237.4822.1601] [PubMed: https://www.ncbi.nlm.nih.gov/pubmed/17834449]

107. Raskin, I. Role of salicylic acid in plants. Annu. Rev. Plant Physiol. Plant Mol. Biol.; 1992; 43, pp. 439-463. [DOI: https://dx.doi.org/10.1146/annurev.pp.43.060192.002255]

108. Khan, M.I.R.; Fatma, M.; Per, T.S.; Anjum, N.A.; Khan, N.A. Salicylic acid-induced abiotic stress tolerance and underlying mechanisms in plants. Front. Plant Sci.; 2015; 6, 462. [DOI: https://dx.doi.org/10.3389/fpls.2015.00462]

109. Miura, K.; Tada, Y. Regulation of water, salinity, and cold stress responses by salicylic acid. Front. Plant Sci.; 2014; 5, 4. [DOI: https://dx.doi.org/10.3389/fpls.2014.00004] [PubMed: https://www.ncbi.nlm.nih.gov/pubmed/24478784]

110. Larkindale, J.; Hall, J.D.; Knight, M.R.; Vierling, E. Heat stress phenotypes of Arabidopsis mutants implicate multiple signaling pathways in the acquisition of thermotolerance. Plant Physiol.; 2005; 138, pp. 882-897. [DOI: https://dx.doi.org/10.1104/pp.105.062257]

111. Rivas-San Vicente, M.; Plasencia, J. Salicylic acid beyond defence: Its role in plant growth and development. J. Exp. Bot.; 2011; 62, pp. 3321-3338. [DOI: https://dx.doi.org/10.1093/jxb/err031]

112. Pasternak, T.; Groot, E.P.; Kazantsev, F.V.; Teale, W.; Omelyanchuk, N.; Kovrizhnykh, V.; Palme, K.; Mironova, V.V. Salicylic acid affects root meristem patterning via auxin distribution in a concentration-dependent manner. Plant Physiol.; 2019; 180, pp. 1725-1739. [DOI: https://dx.doi.org/10.1104/pp.19.00130] [PubMed: https://www.ncbi.nlm.nih.gov/pubmed/31036755]

113. Lee, S.; Kim, S.; Park, C. Salicylic acid promotes seed germination under high salinity by modulating antioxidant activity in Arabidopsis. New Phytol.; 2010; 188, pp. 626-637. [DOI: https://dx.doi.org/10.1111/j.1469-8137.2010.03378.x] [PubMed: https://www.ncbi.nlm.nih.gov/pubmed/20663063]

114. Wildermuth, M.C.; Dewdney, J.; Wu, G.; Ausubel, F.M. Isochorismate synthase is required to synthesize salicylic acid for plant defence. Nature; 2001; 414, pp. 562-565. [DOI: https://dx.doi.org/10.1038/35107108] [PubMed: https://www.ncbi.nlm.nih.gov/pubmed/11734859]

115. Rekhter, D.; Lüdke, D.; Ding, Y.; Feussner, K.; Zienkiewicz, K.; Lipka, V.; Wiermer, M.; Zhang, Y.; Feussner, I. Isochorismate-derived biosynthesis of the plant stress hormone salicylic acid. Science; 2019; 365, pp. 498-502. [DOI: https://dx.doi.org/10.1126/science.aaw1720] [PubMed: https://www.ncbi.nlm.nih.gov/pubmed/31371615]

116. Torrens-Spence, M.P.; Bobokalonova, A.; Carballo, V.; Glinkerman, C.M.; Pluskal, T.; Shen, A.; Weng, J.-K. PBS3 and EPS1 complete salicylic acid biosynthesis from isochorismate in Arabidopsis. Mol. Plant; 2019; 12, pp. 1577-1586. [DOI: https://dx.doi.org/10.1016/j.molp.2019.11.005] [PubMed: https://www.ncbi.nlm.nih.gov/pubmed/31760159]

117. Ullah, C.; Chen, Y.H.; Ortega, M.A.; Tsai, C.J. The diversity of salicylic acid biosynthesis and defense signaling in plants: Knowledge gaps and future opportunities. Curr. Opin. Plant Biol.; 2023; 72, 102349. [DOI: https://dx.doi.org/10.1016/j.pbi.2023.102349] [PubMed: https://www.ncbi.nlm.nih.gov/pubmed/36842224]

118. Yuan, Y.; Chung, J.-D.; Fu, X.; Johnson, V.E.; Ranjan, P.; Booth, S.L.; Harding, S.A.; Tsai, C.-J. Alternative splicing and gene duplication differentially shaped the regulation of isochorismate synthase in Populus and Arabidopsis. Proc. Natl. Acad. Sci. USA; 2009; 106, pp. 22020-22025. [DOI: https://dx.doi.org/10.1073/pnas.0906869106]

119. Holland, C.K.; Westfall, C.S.; Schaffer, J.E.; De Santiago, A.; Zubieta, C.; Alvarez, S.; Jez, J.M. Brassicaceae-specific Gretchen Hagen 3 acyl acid amido synthetases conjugate amino acids to chorismate, a precursor of aromatic amino acids and sali cylic acid. J. Biol. Chem.; 2019; 294, pp. 16855-16864. [DOI: https://dx.doi.org/10.1074/jbc.RA119.009949] [PubMed: https://www.ncbi.nlm.nih.gov/pubmed/31575658]

120. Tada, Y.; Spoel, S.H.; Pajerowska-Mukhtar, K.; Mou, Z.; Song, J.; Wang, C.; Zuo, J.; Dong, X. Plant immunity requires conformational changes of NPR1 via S-nitrosylation and thioredoxins. Science; 2008; 321, pp. 952-956. [DOI: https://dx.doi.org/10.1126/science.1156970] [PubMed: https://www.ncbi.nlm.nih.gov/pubmed/18635760]

121. Bigeard, J.; Colcombet, J.; Hirt, H. Signaling mechanisms in pattern-triggered immunity (PTI). Mol. Plant; 2015; 8, pp. 521-539. [DOI: https://dx.doi.org/10.1016/j.molp.2014.12.022] [PubMed: https://www.ncbi.nlm.nih.gov/pubmed/25744358]

122. Yan, S.; Dong, X. Perception of the plant immune signal salicylic acid. Curr. Opin. Plant Biol.; 2014; 20, pp. 64-68. [DOI: https://dx.doi.org/10.1016/j.pbi.2014.04.006] [PubMed: https://www.ncbi.nlm.nih.gov/pubmed/24840293]

123. Rochon, A.; Boyle, P.; Wignes, T.; Fobert, P.R.; Després, C. The coactivator function of Arabidopsis NPR1 requires the core of its BTB/POZ domain and the oxidation of C-terminal cysteines. Plant Cell; 2006; 18, pp. 3670-3685. [DOI: https://dx.doi.org/10.1105/tpc.106.046953]

124. Boyle, P.; Le Su, E.; Rochon, A.; Shearer, H.L.; Murmu, J.; Chu, J.Y.; Fobert, P.R.; Després, C. The BTB/POZ domain of the Arabidopsis disease resistance protein NPR1 interacts with the repression domain of TGA2 to negate its function. Plant Cell; 2009; 21, pp. 3700-3713. [DOI: https://dx.doi.org/10.1105/tpc.109.069971]

125. Mou, Z.; Fan, W.; Dong, X. Inducers of plant systemic acquired resistance regulate NPR1 function through redox changes. Cell; 2003; 113, pp. 935-944. [DOI: https://dx.doi.org/10.1016/S0092-8674(03)00429-X] [PubMed: https://www.ncbi.nlm.nih.gov/pubmed/12837250]

126. Yuan, Y.; Zhong, S.; Li, Q.; Zhu, Z.; Lou, Y.; Wang, L.; Wang, J.; Wang, M.; Li, Q.; Yang, D. et al. Functional analysis of rice NPR1-like genes reveals that OsNPR1/NH1 is the rice orthologue conferring disease resistance with enhanced herbivore susceptibility. Plant Biotechnol. J.; 2007; 5, pp. 313-324. [DOI: https://dx.doi.org/10.1111/j.1467-7652.2007.00243.x]

127. Jiang, Z.; Jin, X.; Yang, M.; Pi, Q.; Cao, Q.; Li, Z.; Zhang, Y.; Wang, X.-B.; Han, C.; Yu, J. et al. Barley stripe mosaic virus γb protein targets thioredoxin h-type 1 to dampen salicylic acid-mediated defenses. Plant Physiol.; 2022; 189, pp. 1715-1727. [DOI: https://dx.doi.org/10.1093/plphys/kiac137] [PubMed: https://www.ncbi.nlm.nih.gov/pubmed/35325212]

128. Spoel, S.H.; Mou, Z.; Tada, Y.; Spivey, N.W.; Genschik, P.; Dong, X. Proteasome-mediated turnover of the transcription coactivator NPR1 plays dual roles in regulating plant immunity. Cell; 2009; 137, pp. 860-872. [DOI: https://dx.doi.org/10.1016/j.cell.2009.03.038]

129. Fu, Z.Q.; Yan, S.; Saleh, A.; Wang, W.; Ruble, J.; Oka, N.; Mohan, R.; Spoel, S.H.; Tada, Y.; Zheng, N. et al. NPR3 and NPR4 are receptors for the immune signal salicylic acid in plants. Nature; 2012; 486, pp. 228-232. [DOI: https://dx.doi.org/10.1038/nature11162] [PubMed: https://www.ncbi.nlm.nih.gov/pubmed/22699612]

130. Seyfferth, C.; Tsuda, K. Salicylic acid signal transduction: The initiation of biosynthesis, perception and transcriptional reprogramming. Front. Plant Sci.; 2014; 5, 697. [DOI: https://dx.doi.org/10.3389/fpls.2014.00697] [PubMed: https://www.ncbi.nlm.nih.gov/pubmed/25538725]

131. Van Loon, L.C.; Pierpont, W.S.; Boller, T.; Conejero, V. Recommendations for naming plant pathogenesis-related proteins. Plant Mol. Biol. Rep.; 1994; 12, pp. 245-264. [DOI: https://dx.doi.org/10.1007/BF02668748]

132. Okushima, Y.; Koizumi, N.; Kusano, T.; Sano, H. Secreted proteins of tobacco cultured BY2 cells: Identification of a new member of pathogenesis-related proteins. Plant Mol. Biol.; 2000; 42, pp. 479-488. [DOI: https://dx.doi.org/10.1023/A:1006393326985]

133. Edreva, A. Pathogenesis-related proteins: Research progress in the last 15 years. Gen. Appl. Plant Physiol.; 2005; 31, pp. 105-124.

134. Sinha, M.; Singh, R.P.; Kushwaha, G.S.; Iqbal, N.; Singh, A.; Kaushik, S.; Kaur, P.; Sharma, S.; Singh, T.P. Current overview of allergens of plant pathogenesis related protein families. Sci. World J.; 2014; 2014, 543195. [DOI: https://dx.doi.org/10.1155/2014/543195] [PubMed: https://www.ncbi.nlm.nih.gov/pubmed/24696647]

135. Ali, S.; Ganai, B.A.; Kamili, A.N.; Bhat, A.A.; Mir, Z.A.; Bhat, J.A.; Tyagi, A.; Islam, S.T.; Mushtaq, M.; Yadav, P. et al. Pathogenesis-related proteins and peptides as promising tools for engineering plants with multiple stress tolerance. Microbiol. Res.; 2018; 212–213, pp. 29-37. [DOI: https://dx.doi.org/10.1016/j.micres.2018.04.008] [PubMed: https://www.ncbi.nlm.nih.gov/pubmed/29853166]

136. Guevara-Morato, M.A.; de Lacoba, M.G.; García-Luque, I.; Serra, M.T. Characterization of a pathogenesis-related protein 4 (PR-4) induced in Capsicum chinense L3 plants with dual RNase and DNase activities. J. Exp. Bot.; 2010; 61, pp. 3259-3271. [DOI: https://dx.doi.org/10.1093/jxb/erq148] [PubMed: https://www.ncbi.nlm.nih.gov/pubmed/20511278]

137. Ullah, C.; Tsai, C.-J.; Unsicker, S.B.; Xue, L.; Reichelt, M.; Gershenzon, J.; Hammerbacher, A. Salicylic acid activates poplar defense against the biotrophic rust fungus Melampsora larici-populina via increased biosynthesis of catechin and proanthocyanidins. New Phytol.; 2019; 221, pp. 960-975. [DOI: https://dx.doi.org/10.1111/nph.15396] [PubMed: https://www.ncbi.nlm.nih.gov/pubmed/30168132]

138. Gatz, C. From pioneers to team players: TGA transcription factors provide a molecular link between different stress pathways. Mol. Plant Microbe Interact.; 2013; 26, pp. 151-159. [DOI: https://dx.doi.org/10.1094/MPMI-04-12-0078-IA]

139. Wang, D.; Amornsiripanitch, N.; Dong, X. A genomic approach to identify regulatory nodes in the transcriptional network of systemic acquired resistance in plants. PLoS Pathog.; 2006; 2, e123. [DOI: https://dx.doi.org/10.1371/journal.ppat.0020123] [PubMed: https://www.ncbi.nlm.nih.gov/pubmed/17096590]

140. Pajerowska-Mukhtar, K.M.; Wang, W.; Tada, Y.; Oka, N.; Tucker, C.L.; Fonseca, J.P.; Dong, X. The HSF-like transcription factor TBF1 is a major molecular switch for plant growth-to-defense transition. Curr. Biol.; 2012; 22, pp. 103-112. [DOI: https://dx.doi.org/10.1016/j.cub.2011.12.015] [PubMed: https://www.ncbi.nlm.nih.gov/pubmed/22244999]

141. Norman, C.; Howell, K.A.; Millar, A.H.; Whelan, J.M.; Day, D.A. Salicylic acid is an uncoupler and inhibitor of mitochondrial electron transport. Plant Physiol.; 2004; 134, pp. 492-501. [DOI: https://dx.doi.org/10.1104/pp.103.031039] [PubMed: https://www.ncbi.nlm.nih.gov/pubmed/14684840]

142. Chen, Z.; Ricigliano, J.W.; Klessig, D.F. Purification and characterization of a soluble salicylic acid-binding protein from tobacco. Proc. Natl. Acad. Sci. USA; 1993; 90, pp. 9533-9537. [DOI: https://dx.doi.org/10.1073/pnas.90.20.9533] [PubMed: https://www.ncbi.nlm.nih.gov/pubmed/8415736]

143. Durner, J.; Klessig, D.F. Inhibition of ascorbate peroxidase by salicylic acid and 2,6-dichloroisonicotinic acid, two inducers of plant defense responses. Proc. Natl. Acad. Sci. USA; 1995; 92, pp. 11312-11316. [DOI: https://dx.doi.org/10.1073/pnas.92.24.11312] [PubMed: https://www.ncbi.nlm.nih.gov/pubmed/7479986]

144. Jardim-Messeder, D.; Caverzan, A.; Rauber, R.; de Souza Ferreira, E.; Margis-Pinheiro, M.; Galina, A. Succinate dehydrogenase (mitochondrial complex II) is a source of reactive oxygen species in plants and regulates development and stress responses. New Phytol.; 2015; 208, pp. 776-789. [DOI: https://dx.doi.org/10.1111/nph.13515]

145. Belt, K.; Huang, S.; Thatcher, L.F.; Casarotto, H.; Singh, K.B.; Van Aken, O.; Millar, A.H. Salicylic acid-dependent plant stress signaling via mitochondrial succinate dehydrogenase. Plant Physiol.; 2017; 173, pp. 2029-2040. [DOI: https://dx.doi.org/10.1104/pp.16.00060] [PubMed: https://www.ncbi.nlm.nih.gov/pubmed/28209841]

146. Jardim-Messeder, D.; Margis-Pinheiro, M.; Sachetto-Martins, G. Salicylic acid and adenine nucleotides regulate the electron transport system and ROS production in plant mitochondria. Biochim. Biophys. Acta Bioenerg.; 2022; 1863, 148559. [DOI: https://dx.doi.org/10.1016/j.bbabio.2022.148559] [PubMed: https://www.ncbi.nlm.nih.gov/pubmed/35413247]

147. Gleason, C.; Huang, S.; Thatcher, L.F.; Foley, R.C.; Anderson, C.R.; Carroll, A.J.; Millar, A.H.; Singh, K.B. Mitochondrial complex II has a key role in mitochondrial-derived reactive oxygen species influence on plant stress gene regulation and defense. Proc. Natl. Acad. Sci. USA; 2011; 108, pp. 10768-10773. [DOI: https://dx.doi.org/10.1073/pnas.1016060108] [PubMed: https://www.ncbi.nlm.nih.gov/pubmed/21670306]

148. La, V.H.; Lee, B.R.; Islam, M.T.; Park, S.H.; Jung, H.I.; Bae, D.W.; Kim, T.H. Characterization of salicylic acid-mediated modulation of the drought stress responses: Reactive oxygen species, proline, and redox state in Brassica napus. Environ. Exp. Bot.; 2019; 157, pp. 1-10. [DOI: https://dx.doi.org/10.1016/j.envexpbot.2018.09.013]

149. Salem, M.A.; Yoshida, T.; de Souza, L.P.; Alseekh, S.; Bajdzienko, K.; Fernie, A.R.; Giavalisco, P. An improved extraction method enables the com prehensive analysis of lipids, proteins, metabolites and phytohormones from a single sample of leaf tissue under water-defcit stress. Plant J.; 2020; 103, pp. 1614-1632. [DOI: https://dx.doi.org/10.1111/tpj.14800]

150. Lee, J.S. The mechanism of stomatal closing by salicylic acid in Commelina communis L. J. Plant Biol.; 1998; 41, pp. 97-102. [DOI: https://dx.doi.org/10.1007/BF03030395]

151. Khokon, A.R.; Okuma, E.; Hossain, M.A.; Munemasa, S.; Uraji, M.; Nakamura, Y.; Mori, I.C.; Murata, Y. Involvement of extracellular oxidative burst in salicylic acid induced stomatal closure in Arabidopsis. Plant Cell Environ.; 2011; 34, pp. 434-443. [DOI: https://dx.doi.org/10.1111/j.1365-3040.2010.02253.x] [PubMed: https://www.ncbi.nlm.nih.gov/pubmed/21062318]

152. Ananieva, B.A.; Alexieva, V.S.; Popova, L.P. Treatment with salicylic acid decreases the effects of paraquat on photosynthesis. J. Plant Physiol.; 2002; 159, pp. 685-693. [DOI: https://dx.doi.org/10.1078/0176-1617-0706]

153. Loutfy, N.; El-Tayeb, M.A.; Hassanen, A.M.; Moustafa, M.F.M.; Sakuma, Y.; Inouhe, M. Changes in the water status and osmotic solute contents in response to drought and salicylic acid treatments in four different cultivars of wheat (Triticum aestivum) J. Plant Res.; 2012; 125, pp. 173-184. [DOI: https://dx.doi.org/10.1007/s10265-011-0419-9]

154. Mori, I.C.; Pinontoan, R.; Kawano, T.; Muto, S. Involvement of superoxide generation in salicylic acid-induced stomatal closure in Vicia faba. Plant Cell Physiol.; 2001; 42, pp. 1383-1388. [DOI: https://dx.doi.org/10.1093/pcp/pce176]

155. He, J.; Zhang, R.X.; Kim, D.S.; Sun, P.; Liu, H.; Liu, Z.; Hetherington, A.M.; Liang, Y.K. ROS of Distinct Sources and Salicylic Acid Separate Elevated CO2-Mediated Stomatal Movements in Arabidopsis. Front. Plant Sci.; 2020; 11, 542. [DOI: https://dx.doi.org/10.3389/fpls.2020.00542] [PubMed: https://www.ncbi.nlm.nih.gov/pubmed/32457781]

156. Bindschedler, L.V.; Dewdney, J.; Blee, K.A.; Stone, J.M.; Asai, T.; Plotnikov, J.; Denoux, C.; Hayes, T.; Gerrish, C.; Davies, D.R. et al. Peroxidase-dependent apoplastic oxidative burst in Arabidopsis required for pathogen resistance. Plant J.; 2006; 47, pp. 851-863. [DOI: https://dx.doi.org/10.1111/j.1365-313X.2006.02837.x] [PubMed: https://www.ncbi.nlm.nih.gov/pubmed/16889645]

157. Daudi, A.; Cheng, Z.; O’Brien, J.A.; Mammarella, N.; Khan, S.; Ausubel, F.M.; Bolwell, G.P. The apoplastic oxidative burst peroxidase in Arabidopsis is a major component of pattern-triggered immunity. Plant Cell; 2012; 24, pp. 275-287. [DOI: https://dx.doi.org/10.1105/tpc.111.093039] [PubMed: https://www.ncbi.nlm.nih.gov/pubmed/22247251]

158. O’Brien, J.A.; Daudi, A.; Butt, V.S.; Bolwell, G.P. Reactive oxygen species and their role in plant defence and cell wall metabolism. Planta; 2012; 236, pp. 765-779. [DOI: https://dx.doi.org/10.1007/s00425-012-1696-9] [PubMed: https://www.ncbi.nlm.nih.gov/pubmed/22767200]

159. O’Brien, J.A.; Daudi, A.; Finch, P.; Butt, V.S.; Whitelegge, J.P.; Souda, P.; Ausubel, F.M.; Bolwell, G.P. A peroxidase-dependent apoplastic oxidative burst in cultured Arabidopsis cells functions in MAMP-elicited defense. Plant Physiol.; 2012; 158, pp. 2013-2027. [DOI: https://dx.doi.org/10.1104/pp.111.190140]

160. Arnaud, D.; Lee, S.; Takebayashi, Y.; Choi, D.; Choi, J.; Sakakibara, H.; Hwang, I. Cytokinin-Mediated Regulation of Reactive Oxygen Species Homeostasis Modulates Stomatal Immunity in Arabidopsis. Plant Cell; 2017; 29, pp. 543-559. [DOI: https://dx.doi.org/10.1105/tpc.16.00583]

161. Saito, S.; Uozumi, N. Guard Cell Membrane Anion Transport Systems and Their Regulatory Components: An Elaborate Mechanism Controlling Stress-Induced Stomatal Closure. Plants; 2019; 8, 9. [DOI: https://dx.doi.org/10.3390/plants8010009] [PubMed: https://www.ncbi.nlm.nih.gov/pubmed/30609843]

162. Zeng, W.; He, S.Y. A prominent role of the fagellin receptor FLAGELLIN SENSING2 in mediating stomatal response to Pseudomonas syringae pv tomato DC3000 in Arabidopsis. Plant Physiol.; 2010; 153, pp. 1188-1198. [DOI: https://dx.doi.org/10.1104/pp.110.157016]

163. Wang, H.Q.; Sun, L.P.; Wang, L.X.; Fang, X.W.; Li, Z.Q.; Zhang, F.F.; Hu, X.; Qi, C.; He, J.M. Ethylene mediates salicylic acid induced stomatal closure by controlling reactive oxygen species and nitric oxide production in Arabidopsis. Plant Sci.; 2020; 294, 110464. [DOI: https://dx.doi.org/10.1016/j.plantsci.2020.110464]

164. Mao, Y.B.; Liu, Y.Q.; Chen, D.Y.; Chen, F.Y.; Fang, X.; Hong, G.J.; Wang, L.J.; Wang, J.W.; Chen, X.Y. Jasmonate response decay and defense metabolite accumulation contributes to age-regulated dynamics of plant insect resistance. Nat. Commun.; 2017; 8, 13925. [DOI: https://dx.doi.org/10.1038/ncomms13925]

165. Verma, V.; Ravindran, P.; Kumar, P.P. Plant hormone-mediated regulation of stress responses. BMC Plant Biol.; 2016; 16, 86. [DOI: https://dx.doi.org/10.1186/s12870-016-0771-y] [PubMed: https://www.ncbi.nlm.nih.gov/pubmed/27079791]

166. Yang, J.; Duan, G.; Li, C.; Liu, L.; Han, G.; Zhang, Y.; Wang, C. The crosstalks between jasmonic acid and other plant hormone signaling highlight the involvement of jasmonic acid as a core component in plant response to biotic and abiotic stresses. Front. Plant Sci.; 2019; 10, 1349. [DOI: https://dx.doi.org/10.3389/fpls.2019.01349]

167. Zander, M.; Lewsey, M.G.; Clark, N.M.; Yin, L.; Bartlett, A.; Saldierna Guzmán, J.P.; Hann, E.; Langford, A.E.; Jow, B.; Wise, A. et al. Integrated multi-omics framework of the plant response to jasmonic acid. Nat. Plants; 2020; 6, pp. 290-302. [DOI: https://dx.doi.org/10.1038/s41477-020-0605-7] [PubMed: https://www.ncbi.nlm.nih.gov/pubmed/32170290]

168. Browse, J. Jasmonate passes muster: A receptor and targets for the defense hormone. Annu. Rev. Plant Biol.; 2009; 60, pp. 183-205. [DOI: https://dx.doi.org/10.1146/annurev.arplant.043008.092007] [PubMed: https://www.ncbi.nlm.nih.gov/pubmed/19025383]

169. Fonseca, S.; Chini, A.; Hamberg, M.; Adie, B.; Porzel, A.; Kramell, R.; Miersch, O.; Wasternack, C.; Solano, R. (+)-7-iso-Jasmonoyl-L-isoleucine is the endogenous bioactive jasmonate. Nat. Chem. Biol.; 2009; 5, pp. 344-350. [DOI: https://dx.doi.org/10.1038/nchembio.161]

170. Gfeller, A.; Dubugnon, L.; Liechti, R.; Farmer, E.E. Jasmonate biochemical pathway. Sci. Signal.; 2010; 3, cm3. [PubMed: https://www.ncbi.nlm.nih.gov/pubmed/20159849][DOI: https://dx.doi.org/10.1126/scisignal.3109cm3]

171. Mahmud, S.; Ullah, C.; Kortz, A.; Bhattacharyya, S.; Yu, P.; Gershenzon, J.; Vothknecht, U.C. Constitutive expression of JASMONATE RESISTANT 1 induces molecular changes that prime the plants to better withstand drought. Plant Cell Environ.; 2022; 45, pp. 2906-2922. [DOI: https://dx.doi.org/10.1111/pce.14402]

172. Koo, A.J.; Cooke, T.F.; Howe, G.A. Cytochrome P450 CYP94B3 mediates catabolism and inactivation of the plant hormone jasmonoyl-L-isoleucine. Proc. Natl. Acad. Sci. USA; 2011; 108, pp. 9298-9303.PMC3107288[DOI: https://dx.doi.org/10.1073/pnas.1103542108] [PubMed: https://www.ncbi.nlm.nih.gov/pubmed/21576464]

173. Li, Q.; Zheng, J.; Li, S.; Huang, G.; Skilling, S.J.; Wang, L.; Li, L.; Li, M.; Yuan, L.; Liu, P. Transporter-Mediated Nuclear Entry of Jasmonoyl-Isoleucine Is Essential for Jasmonate Signaling. Mol. Plant; 2017; 10, pp. 695-708. [DOI: https://dx.doi.org/10.1016/j.molp.2017.01.010]

174. Turner, J.G.; Ellis, C.; Devoto, A. The jasmonate signal pathway. Plant Cell; 2002; 14, pp. S153-S164. [DOI: https://dx.doi.org/10.1105/tpc.000679]

175. Yan, J.; Yao, R.; Chen, L.; Li, S.; Gu, M.; Nan, F.; Xie, D. Dynamic Perception of Jasmonates by the F-Box Protein COI1. Mol. Plant; 2018; 11, pp. 1237-1247. [DOI: https://dx.doi.org/10.1016/j.molp.2018.07.007]

176. Ke, J.; Ma, H.; Gu, X.; Thelen, A.; Brunzelle, J.S.; Li, J.; Xu, H.E.; Melcher, K. Structural basis for recognition of diverse transcriptional repressors by the TOPLESS family of corepressors. Sci. Adv.; 2015; 1, e1500107. [DOI: https://dx.doi.org/10.1126/sciadv.1500107] [PubMed: https://www.ncbi.nlm.nih.gov/pubmed/26601214]

177. Ali, M.S.; Baek, K.H. Jasmonic acid signaling pathway in response to abiotic stresses in plants. Int. J. Mol. Sci.; 2020; 21, 621. [DOI: https://dx.doi.org/10.3390/ijms21020621] [PubMed: https://www.ncbi.nlm.nih.gov/pubmed/31963549]

178. Bäckström, S.; Elfving, N.; Nilsson, R.; Wingsle, G.; Björklund, S. Purification of a plant mediator from Arabidopsis thaliana identifies PFT1 as the Med25 subunit. Mol. Cell; 2007; 26, pp. 717-729. [DOI: https://dx.doi.org/10.1016/j.molcel.2007.05.007]

179. Chen, R.; Jiang, H.; Li, L.; Zhai, Q.; Qi, L.; Zhou, W.; Liu, X.; Li, H.; Zheng, W.; Sun, J. et al. The Arabidopsis mediator subunit MED25 differentially regulates jasmonate and abscisic acid signaling through interacting with the MYC2 and ABI5 transcription factors. Plant Cell; 2012; 24, pp. 2898-2916. [DOI: https://dx.doi.org/10.1105/tpc.112.098277] [PubMed: https://www.ncbi.nlm.nih.gov/pubmed/22822206]

180. Chini, A.; Gimenez-Ibanez, S.; Goossens, A.; Solano, R. Redundancy and specificity in jasmonate signalling. Curr. Opin. Plant Biol.; 2016; 33, pp. 147-156. [DOI: https://dx.doi.org/10.1016/j.pbi.2016.07.005]

181. Wasternack, C.; Song, S. Jasmonates: Biosynthesis, metabolism, and signaling by proteins activating and repressing transcription. J. Exp. Bot.; 2017; 68, pp. 1303-1321. [DOI: https://dx.doi.org/10.1093/jxb/erw443] [PubMed: https://www.ncbi.nlm.nih.gov/pubmed/27940470]

182. Pauwels, L.; Barbero, G.F.; Geerinck, J.; Tilleman, S.; Grunewald, W.; Pérez, A.C.; Chico, J.M.; Bossche, R.V.; Sewell, J.; Gil, E. et al. NINJA connects the co-repressor TOPLESS to jasmonate signalling. Nature; 2010; 464, pp. 788-791. [DOI: https://dx.doi.org/10.1038/nature08854]

183. Chen, H.; Gonzales-Vigil, E.; Wilkerson, C.G.; Howe, G.A. Stability of plant defense proteins in the gut of insect herbivores. Plant Physiol.; 2007; 143, pp. 1954-1967. [DOI: https://dx.doi.org/10.1104/pp.107.095588] [PubMed: https://www.ncbi.nlm.nih.gov/pubmed/17416643]

184. Tan, C.W.; Chiang, S.Y.; Ravuiwasa, K.T.; Yadav, J.; Hwang, S.Y. Jasmonate-induced defenses in tomato against Helicoverpa armigera depend in part on nutrient availability, but artificial induction via methyl jasmonate does not. Arthropod. Plant Interact.; 2012; 6, pp. 531-541. [DOI: https://dx.doi.org/10.1007/s11829-012-9206-3]

185. Yan, J.; Tan, Y.; Lv, Y.; Wang, F.; Zhang, Y.; Chi, D. Effect of jasmonate treatments on leaves of Rosa rugosa ‘Plena’ and detoxification enzymes and feeding of adult Monolepta hieroglyphica. J. For. Res.; 2021; 32, pp. 1253-1261. [DOI: https://dx.doi.org/10.1007/s11676-020-01278-5]

186. Li, R.; Jin, J.; Xu, J.; Wang, L.; Li, J.; Lou, Y.; Baldwin, I.T. Long non-coding RNAs associate with jasmonate-mediated plant defence against herbivores. Plant Cell Environ.; 2021; 44, pp. 982-994. [DOI: https://dx.doi.org/10.1111/pce.13952]

187. Maruta, T.; Inoue, T.; Tamoi, M.; Yabuta, Y.; Yoshimura, K.; Ishikawa, T.; Shigeoka, S. Arabidopsis NADPH oxidases, AtrbohD and AtrbohF, are essential for jasmonic acid-induced expression of genes regulated by MYC2 transcription factor. Plant Sci.; 2011; 180, pp. 655-660. [DOI: https://dx.doi.org/10.1016/j.plantsci.2011.01.014] [PubMed: https://www.ncbi.nlm.nih.gov/pubmed/21421415]

188. Orozco-Cárdenas, M.L.; Narváez-Vásquez, J.; Ryan, C.A. Hydrogen peroxide acts as a second messenger for the induction of defense genes in tomato plants in response to wounding, systemin, and methyl jasmonate. Plant Cell; 2001; 13, pp. 179-191. [DOI: https://dx.doi.org/10.1105/tpc.13.1.179] [PubMed: https://www.ncbi.nlm.nih.gov/pubmed/11158538]

189. Huang, X.; Stettmaier, K.; Michel, C.; Hutzler, P.; Mueller, M.J.; Durner, J. Nitric oxide is induced by wounding and influences jasmonic acid signaling in Arabidopsis thaliana. Planta; 2004; 218, pp. 938-946. [DOI: https://dx.doi.org/10.1007/s00425-003-1178-1]

190. Orozco-Cárdenas, M.; Ryan, C.A. Nitric oxide negatively modulates wound signaling in tomato plants. Plant Physiol.; 2002; 130, pp. 487-493. [DOI: https://dx.doi.org/10.1104/pp.008375] [PubMed: https://www.ncbi.nlm.nih.gov/pubmed/12226527]

191. Hung, K.T.; Kao, C.H. Nitric oxide acts as an antioxidant and delays methyl jasmonate-induced senescence of rice leaves. J. Plant Physiol.; 2004; 161, pp. 43-52. [DOI: https://dx.doi.org/10.1078/0176-1617-01178] [PubMed: https://www.ncbi.nlm.nih.gov/pubmed/15002663]

192. Yang, Y.; Ahammed, G.; Wu, C.; Fan, S.; Zhou, Y. Crosstalk among jasmonate, salicylate and ethylene signaling pathways in plant disease and immune responses. Curr. Protein Pept. Sci.; 2015; 16, pp. 450-461. [DOI: https://dx.doi.org/10.2174/1389203716666150330141638]

193. Howe, G.A.; Major, I.T.; Koo, A.J. Modularity in jasmonate signaling for multistress resilience. Annu. Rev. Plant Biol.; 2018; 69, pp. 387-415. [DOI: https://dx.doi.org/10.1146/annurev-arplant-042817-040047] [PubMed: https://www.ncbi.nlm.nih.gov/pubmed/29539269]

194. Nomoto, M.; Skelly, M.J.; Itaya, T.; Mori, T.; Suzuki, T.; Matsushita, T.; Tokizawa, M.; Kuwata, K.; Mori, H.; Yamamoto, Y.Y. et al. Suppression of MYC transcription activators by the immune cofactor NPR1 fine-tunes plant immune responses. Cell Rep.; 2021; 37, 110125. [DOI: https://dx.doi.org/10.1016/j.celrep.2021.110125]

195. Zheng, X.Y.; Spivey, N.W.; Zeng, W.; Liu, P.P.; Fu, Z.Q.; Klessig, D.F.; He, S.Y.; Dong, X. Coronatine promotes pseudomonas syringae virulence in plants by activating a signaling cascade that inhibits salicylic acid accumulation. Cell Host Microbe; 2012; 11, pp. 587-596. [DOI: https://dx.doi.org/10.1016/j.chom.2012.04.014]

196. Wasternack, C.; Hause, B. Jasmonates: Biosynthesis, perception, signal transduction and action in plant stress response, growth and development. An update to the 2007 review in Annals of Botany. Ann. Bot.; 2013; 111, pp. 1021-1058. [DOI: https://dx.doi.org/10.1093/aob/mct067] [PubMed: https://www.ncbi.nlm.nih.gov/pubmed/23558912]

197. Song, C.; Cao, Y.; Dai, J.; Li, G.; Manzoor, M.A.; Chen, C.; Deng, H. The Multifaceted Roles of MYC2 in Plants: Toward Transcriptional Reprogramming and Stress Tolerance by Jasmonate Signaling. Front. Plant Sci.; 2022; 13, 868874. [DOI: https://dx.doi.org/10.3389/fpls.2022.868874] [PubMed: https://www.ncbi.nlm.nih.gov/pubmed/35548315]

198. Fu, J.; Wu, H.; Ma, S.; Xiang, D.; Liu, R.; Xiong, L. OsJAZ1 attenuates drought resistance by regulating JA and ABA signaling in rice. Front. Plant Sci.; 2017; 8, 2108. [DOI: https://dx.doi.org/10.3389/fpls.2017.02108] [PubMed: https://www.ncbi.nlm.nih.gov/pubmed/29312378]

199. Riemann, M.; Dhakarey, R.; Hazman, M.; Miro, B.; Kohli, A.; Nick, P. Exploring jasmonates in the hormonal network of drought and salinity responses. Front. Plant Sci.; 2015; 6, 1077. [DOI: https://dx.doi.org/10.3389/fpls.2015.01077] [PubMed: https://www.ncbi.nlm.nih.gov/pubmed/26648959]

200. Fang, L.; Su, L.; Sun, X.; Li, X.; Sun, M.; Karungo, S.K.; Fang, S.; Chu, J.; Li, S.; Xin, H. Expression of Vitis amurensis NAC26 in Arabidopsis enhances drought tolerance by modulating jasmonic acid synthesis. J. Exp. Bot.; 2016; 67, pp. 2829-2845. [DOI: https://dx.doi.org/10.1093/jxb/erw122] [PubMed: https://www.ncbi.nlm.nih.gov/pubmed/27162276]

201. Kazan, K.; Manners, J.M. The interplay between light and jasmonate signalling during defence and development. J. Exp. Bot.; 2011; 62, pp. 4087-4100. [DOI: https://dx.doi.org/10.1093/jxb/err142] [PubMed: https://www.ncbi.nlm.nih.gov/pubmed/21705384]

202. Savchenko, T.; Dehesh, K. Drought stress modulates oxylipin signature by eliciting 12-OPDA as a potent regulator of stomatal aperture. Plant Signal. Behav.; 2014; 9, e28304. [DOI: https://dx.doi.org/10.4161/psb.28304] [PubMed: https://www.ncbi.nlm.nih.gov/pubmed/24614570]

203. Ayyaz, A.; Miao, Y.; Hannan, F.; Islam, F.; Zhang, K.; Xu, J.; Farooq, M.A.; Zhou, W. Drought tolerance in Brassica napus is accompanied with enhanced antioxidative protection, photosynthetic and hormonal regulation at seedling stage. Physiol. Plant.; 2021; 172, pp. 1133-1148. [DOI: https://dx.doi.org/10.1111/ppl.13375] [PubMed: https://www.ncbi.nlm.nih.gov/pubmed/33599291]

204. Ho, T.T.; Murthy, H.N.; Park, S.Y. Methyl Jasmonate Induced Oxidative Stress and Accumulation of Secondary Metabolites in Plant Cell and Organ Cultures. Int. J. Mol. Sci.; 2020; 21, 716. [DOI: https://dx.doi.org/10.3390/ijms21030716]

205. Mittler, R. Oxidative stress, antioxidants and stress tolerance. Trends Plant Sci.; 2002; 7, pp. 405-410. [DOI: https://dx.doi.org/10.1016/S1360-1385(02)02312-9] [PubMed: https://www.ncbi.nlm.nih.gov/pubmed/12234732]

206. Shan, C.; Zhou, Y.; Liu, M. Nitric oxide participates in the regulation of the ascorbate-glutathione cycle by exogenous jasmonic acid in the leaves of wheat seedlings under drought stress. Protoplasma; 2015; 252, pp. 1397-1405. [DOI: https://dx.doi.org/10.1007/s00709-015-0756-y] [PubMed: https://www.ncbi.nlm.nih.gov/pubmed/25577230]

207. Camara, M.C.; Vandenberghe, L.P.S.; Rodrigues, C.; de Oliveira, J.; Faulds, C.; Bertrand, E.; Soccol, C.R. Current advances in gibberellic acid (GA3) production, patented technologies and potential applications. Planta; 2018; 248, pp. 1049-1062. [DOI: https://dx.doi.org/10.1007/s00425-018-2959-x] [PubMed: https://www.ncbi.nlm.nih.gov/pubmed/30069731]

208. Bon, D.J.Y.D.; Mander, L.N.; Lan, P. Syntheses of Gibberellins A15 and A24, the Key Metabolites in Gibberellin Biosynthesis. J. Org. Chem.; 2018; 83, pp. 6566-6572. [DOI: https://dx.doi.org/10.1021/acs.joc.8b00876]

209. Magome, H.; Nomura, T.; Hanada, A.; Takeda-Kamiya, N.; Ohnishi, T.; Shinma, Y.; Katsumata, T.; Kawaide, H.; Kamiya, Y.; Yamaguchi, S. CYP714B1 and CYP714B2 encode gibberellin 13-oxidases that reduce gibberellin activity in rice. Proc. Natl. Acad. Sci. USA; 2013; 110, pp. 1947-1952. [DOI: https://dx.doi.org/10.1073/pnas.1215788110]

210. Rieu, I.; Eriksson, S.; Powers, S.J.; Gong, F.; Griffiths, J.; Woolley, L.; Benlloch, R.; Nilsson, O.; Thomas, S.G.; Hedden, P. et al. Genetic analysis reveals that C19-GA 2-oxidation is a major gibberellin inactivation pathway in Arabidopsis. Plant Cell; 2008; 20, pp. 2420-2436. [DOI: https://dx.doi.org/10.1105/tpc.108.058818]

211. Nir, I.; Moshelion, M.; Weiss, D. The Arabidopsis GIBBERELLIN METHYL TRANSFERASE 1 suppresses gibberellin activity, reduces whole-plant transpiration and promotes drought tolerance in transgenic tomato. Plant Cell Environ.; 2014; 37, pp. 113-123. [DOI: https://dx.doi.org/10.1111/pce.12135]

212. Kanno, Y.; Oikawa, T.; Chiba, Y.; Ishimaru, Y.; Shimizu, T.; Sano, N.; Koshiba, T.; Kamiya, Y.; Ueda, M.; Seo, M. AtSWEET13 and AtSWEET14 regulate gibberellin-mediated physiological processes. Nat. Commun.; 2016; 7, 13245. [DOI: https://dx.doi.org/10.1038/ncomms13245] [PubMed: https://www.ncbi.nlm.nih.gov/pubmed/27782132]

213. Chiba, Y.; Shimizu, T.; Miyakawa, S.; Kanno, Y.; Koshiba, T.; Kamiya, Y.; Seo, M. Identification of Arabidopsis thaliana NRT1/PTR FAMILY (NPF) proteins capable of transporting plant hormones. J. Plant Res.; 2015; 128, pp. 679-686. [DOI: https://dx.doi.org/10.1007/s10265-015-0710-2]

214. Schwechheime, C. Gibberellin signaling in plants—The extended version. Front. Plant Sci.; 2012; 2, 107. [DOI: https://dx.doi.org/10.3389/fpls.2011.00107]

215. Phokas, A.; Coates, J.C. Evolution of DELLA function and signaling in land plants. Evol. Dev.; 2021; 23, pp. 137-154. [DOI: https://dx.doi.org/10.1111/ede.12365] [PubMed: https://www.ncbi.nlm.nih.gov/pubmed/33428269]

216. Huang, X.; Tian, H.; Park, J.; Oh, D.H.; Hu, J.; Zentella, R.; Qiao, H.; Dassanayake, M.; Sun, T.P. The master growth regulator DELLA binding to histone H2A is essential for DELLA-mediated global transcription regulation. Nat. Plants; 2023; 9, pp. 1291-1305. [DOI: https://dx.doi.org/10.1038/s41477-023-01477-y] [PubMed: https://www.ncbi.nlm.nih.gov/pubmed/37537399]

217. Sasaki, A.; Itoh, H.; Gomi, K.; Ueguchi-Tanaka, M.; Ishiyama, K.; Kobayashi, M.; Jeong, D.H.; An, G.; Kitano, H.; Ashikari, M. et al. Accumulation of phosphorylated repressor for gibberellin signaling in an F-box mutant. Science; 2003; 299, pp. 1896-1898. [DOI: https://dx.doi.org/10.1126/science.1081077] [PubMed: https://www.ncbi.nlm.nih.gov/pubmed/12649483]

218. Gomi, K.; Sasaki, A.; Itoh, H.; Ueguchi-Tanaka, M.; Ashikari, M.; Kitano, H.; Matsuoka, M. GID2, an F-box subunit of the SCF E3 complex, specifically interacts with phosphorylated SLR1 protein and regulates the gibberellin-dependent degradation of SLR1 in rice. Plant J.; 2004; 37, pp. 626-634. [DOI: https://dx.doi.org/10.1111/j.1365-313X.2003.01990.x]

219. Mcginnis, K.M.; Thomas, S.G.; Soule, J.D.; Strader, L.C.; Zale, J.M.; Sun, T.P.; Steber, C.M. The Arabidopsis SLEEPY1 gene encodes a putative F-box subunit of an SCF E3 ubiquitin ligase. Plant Cell; 2003; 15, pp. 1120-1130. [DOI: https://dx.doi.org/10.1105/tpc.010827]

220. Dill, A.; Thomas, S.G.; Hu, J.; Steber, C.M.; Sun, T.P. The Arabidopsis F-box protein SLEEPY1 targets gibberellin signaling repressors for gibberellin-induced degradation. Plant Cell; 2004; 16, pp. 1392-1405. [DOI: https://dx.doi.org/10.1105/tpc.020958]

221. Fu, X.; Richards, D.E.; Fleck, B.; Xie, D.; Burton, N.; Harberd, N.P. The Arabidopsis mutant sleepy1gar2-1 protein promotes plant growth by increasing the affinity of the SCFSLY1 E3 ubiquitin ligase for DELLA protein substrates. Plant Cell; 2004; 16, pp. 1406-1418. [DOI: https://dx.doi.org/10.1105/tpc.021386] [PubMed: https://www.ncbi.nlm.nih.gov/pubmed/15161962]

222. Dohmann, E.M.; Nill, C.; Schwechheimer, C. DELLA proteins restrain germination and elongation growth in Arabidopsis thaliana COP9 signalosome mutants. Eur. J. Cell Biol.; 2010; 89, pp. 163-168. [DOI: https://dx.doi.org/10.1016/j.ejcb.2009.12.001] [PubMed: https://www.ncbi.nlm.nih.gov/pubmed/20083325]

223. Ariizumi, T.; Lawrence, P.K.; Steber, C.M. The role of two F-box proteins, SLEEPY1 and SNEEZY, in Arabidopsis gibberellin signaling. Plant Physiol.; 2011; 155, pp. 765-775. [DOI: https://dx.doi.org/10.1104/pp.110.166272] [PubMed: https://www.ncbi.nlm.nih.gov/pubmed/21163960]

224. Ariizumi, T.; Murase, K.; Sun, T.-P.; Steber, C.M. Proteolysis independent down-regulation of DELLA repression by the gibberellin receptor GIBBERELLIN INSENSI TIVE DWARF 1. Plant Cell; 2008; 20, pp. 2447-2459. [DOI: https://dx.doi.org/10.1105/tpc.108.058487] [PubMed: https://www.ncbi.nlm.nih.gov/pubmed/18827182]

225. Ueguchi-Tanaka, M.; Hirano, K.; Hasegawa, Y.; Kitano, H.; Matsuoka, M. Release of the repressive activity of rice DELLA protein SLR1 by gibberellin does not require SLR1 degradation in the gid2 mutant. Plant Cell; 2008; 20, pp. 2437-2446. [DOI: https://dx.doi.org/10.1105/tpc.108.061648] [PubMed: https://www.ncbi.nlm.nih.gov/pubmed/18827181]

226. De Lucas, M.; Daviere, J.M.; Rodriguez-Falcon, M.; Pontin, M.; Iglesias-Pedraz, J.M.; Lorrain, S.; Fankhauser, C.; Blazquez, M.A.; Titarenko, E.; Prat, S. A molecular framework for light and gibberellin control of cell elongation. Nature; 2008; 451, pp. 480-484. [DOI: https://dx.doi.org/10.1038/nature06520] [PubMed: https://www.ncbi.nlm.nih.gov/pubmed/18216857]

227. Feng, S.; Martinez, C.; Gusmaroli, G.; Wang, Y.; Zhou, J.; Wang, F.; Chen, L.; Yu, L.; Iglesias-Pedraz, J.M.; Kircher, S. et al. Coordinated regulation of Arabidopsis thaliana development by light and gibberellins. Nature; 2008; 451, pp. 475-479. [DOI: https://dx.doi.org/10.1038/nature06448]

228. Arnaud, N.; Girin, T.; Sorefan, K.; Fuentes, S.; Wood, T.A.; Lawrenson, T.; Sablowski, R.; Ostergaard, L. Gibberellins control fruit patterning in Arabidopsis thaliana. Genes Dev.; 2010; 24, pp. 2127-2132. [DOI: https://dx.doi.org/10.1101/gad.593410] [PubMed: https://www.ncbi.nlm.nih.gov/pubmed/20889713]

229. Josse, E.M.; Gan, Y.; Bou-Torrent, J.; Stewart, K.L.; Gilday, A.D.; Jeffree, C.E.; Vaistij, F.E.; Martinez-Garcia, J.F.; Nagy, F.; Graham, I.A. et al. A DELLA in disguise: SPATULA restrains the growth of the developing Ara bidopsis seedling. Plant Cell; 2011; 23, pp. 1337-1351. [DOI: https://dx.doi.org/10.1105/tpc.110.082594] [PubMed: https://www.ncbi.nlm.nih.gov/pubmed/21478445]

230. Gubler, F.; Chandler, P.M.; White, R.G.; Llewellyn, D.J.; Jacobsen, J.V. Gibberellin signaling in barley aleurone cells. Control of SLN1 and GAMYB expression. Plant Physiol.; 2002; 129, pp. 191-200. [DOI: https://dx.doi.org/10.1104/pp.010918] [PubMed: https://www.ncbi.nlm.nih.gov/pubmed/12011350]

231. Barba-Espin, G.; Diaz-Vivancos, P.; Clemente-Moreno, M.J.; Albacete, A.; Faize, L.; Faize, M.; Pérez-Alfocea, F.; Hernández, J.A. Interaction between hydrogen peroxide and plant hormones during germination and the early growth of pea seedlings. Plant Cell Environ.; 2010; 33, pp. 981-994. [DOI: https://dx.doi.org/10.1111/j.1365-3040.2010.02120.x] [PubMed: https://www.ncbi.nlm.nih.gov/pubmed/20102539]

232. Fath, A.; Bethke, P.C.; Jones, R.L. Enzymes that scavenge reactive oxygen species are down-regulated prior to gibberellic acid-induced programmed cell death in barley aleurone. Plant Physiol.; 2001; 126, pp. 156-166. [DOI: https://dx.doi.org/10.1104/pp.126.1.156]

233. Bethke, P.C.; Jones, R.L. Cell death of barley aleurone protoplasts is mediated by reactive oxygen species. Plant J.; 2001; 25, pp. 19-29. [DOI: https://dx.doi.org/10.1111/j.1365-313X.2001.00930.x]

234. Ishibashi, Y.; Tawaratsumida, T.; Kondo, K.; Kasa, S.; Sakamoto, M.; Aoki, N.; Zheng, S.H.; Yuasa, T.; Iwaya-Inoue, M. Reactive oxygen species are involved in gibberellin/abscisic acid signaling in barley aleurone cells. Plant Physiol.; 2012; 158, pp. 1705-1714. [DOI: https://dx.doi.org/10.1104/pp.111.192740]

235. Magome, H.; Kamiya, Y. Inactivation processes. Annual Plant Reviews; Hedden, P.; Thomas, S.G. Wiley: Hoboken, NJ, USA, 2016; Volume 49, [DOI: https://dx.doi.org/10.1002/9781119210436.ch3]

236. Tuna, A.L.; Kaya, C.; Dikilitas, M.; Higgs, D. The combined effects of gibberellic acid and salinity on some antioxidant enzyme activities, plant growth parameters and nutritional status in maize plants. Environ. Exp. Bot.; 2008; 62, pp. 1-9. [DOI: https://dx.doi.org/10.1016/j.envexpbot.2007.06.007]

237. Omena Garcia, R.; Martins, A.; Medeiros, D.; Vallarino, J.; Ribeiro, D.; Fernie, A.; Araujo, W.; Nunes, N.A. Growth and metabolic adjustments in response to gibberellin deficiency in drought stressed tomato plants. Environ. Exp. Bot.; 2019; 159, pp. 95-107. [DOI: https://dx.doi.org/10.1016/j.envexpbot.2018.12.011]

238. Achard, P.; Renou, J.P.; Berthomé, R.; Harberd, N.P.; Genschik, P. Plant DELLAs restrain growth and promote survival of adversity by reducing the levels of reactive oxygen species. Curr. Biol.; 2008; 18, pp. 656-660. [DOI: https://dx.doi.org/10.1016/j.cub.2008.04.034] [PubMed: https://www.ncbi.nlm.nih.gov/pubmed/18450450]

239. Colebrook, E.H.; Thomas, S.G.; Phillips, A.L.; Hedden, P. The role of gibberellin signalling in plant responses to abiotic stress. J. Exp. Biol.; 2014; 217, pp. 67-75. [DOI: https://dx.doi.org/10.1242/jeb.089938] [PubMed: https://www.ncbi.nlm.nih.gov/pubmed/24353205]

240. Illouz-Eliaz, N.; Nissan, I.; Nir, I.; Ramon, U.; Shohat, H.; Weiss, D. Mutations in the tomato gibberellin receptors suppress xylem proliferation and reduce water loss under water-deficit conditions. J. Exp. Bot.; 2020; 71, pp. 3603-3612. [DOI: https://dx.doi.org/10.1093/jxb/eraa137] [PubMed: https://www.ncbi.nlm.nih.gov/pubmed/32173726]

241. Illouz-Eliaz, N.; Ramon, U.; Shohat, H.; Blum, S.; Livne, S.; Mendelson, D.; Weiss, D. Multiple gibberellin receptors contribute to phenotypic stability under changing environments. Plant Cell; 2019; 31, pp. 1506-1519. [DOI: https://dx.doi.org/10.1105/tpc.19.00235] [PubMed: https://www.ncbi.nlm.nih.gov/pubmed/31076539]

242. Nir, I.; Shohat, H.; Panizel, I.; Olszewski, N.; Aharoni, A.; Weiss, D. The tomato DELLA protein PROCERA acts in guard cells to promote stomatal closure. Plant Cell; 2017; 29, pp. 3186-3197. [DOI: https://dx.doi.org/10.1105/tpc.17.00542] [PubMed: https://www.ncbi.nlm.nih.gov/pubmed/29150547]

243. Woodward, A.W.; Bartel, B. Auxin: Regulation, action, and interaction. Ann. Bot.; 2005; 95, pp. 707-735. [DOI: https://dx.doi.org/10.1093/aob/mci083]

244. Dai, X.; Mashiguchi, K.; Chen, Q.; Kasahara, H.; Kamiya, Y.; Ojha, S.; DuBois, J.; Ballou, D.; Zhao, Y. The biochemical mechanism of auxin biosynthesis by an arabidopsis YUCCA flavin-containing monooxygenase. J. Biol. Chem.; 2013; 288, pp. 1448-1457. [DOI: https://dx.doi.org/10.1074/jbc.M112.424077]

245. Zhao, Y.; Christensen, S.K.; Fankhauser, C.; Cashman, J.R.; Cohen, J.D.; Weigel, D.; Chory, J. A role for flavin monooxygenase-like enzymes in auxin biosynthesis. Science; 2001; 291, pp. 306-309. [DOI: https://dx.doi.org/10.1126/science.291.5502.306] [PubMed: https://www.ncbi.nlm.nih.gov/pubmed/11209081]

246. Zheng, Z.; Guo, Y.; Novák, O.; Chen, W.; Ljung, K.; Noel, J.P.; Chory, J. Local auxin metabolism regulates environment-induced hypocotyl elongation. Nat. Plants; 2016; 2, 16025. [DOI: https://dx.doi.org/10.1038/nplants.2016.25]

247. Hill, K. Post-translational modifications of hormone-responsive transcription factors: The next level of regulation. J. Exp. Bot.; 2015; 66, pp. 4933-4945. [DOI: https://dx.doi.org/10.1093/jxb/erv273] [PubMed: https://www.ncbi.nlm.nih.gov/pubmed/26041319]

248. Petrasek, J.; Friml, J. Auxin transport routes in plant development. Development; 2009; 136, pp. 2675-2688. [DOI: https://dx.doi.org/10.1242/dev.030353] [PubMed: https://www.ncbi.nlm.nih.gov/pubmed/19633168]

249. Peer, W.A.; Blakeslee, J.J.; Yang, H.; Murphy, A.S. Seven things we think we know about auxin transport. Mol. Plant; 2011; 4, pp. 487-504. [DOI: https://dx.doi.org/10.1093/mp/ssr034]

250. Fukaki, H.; Tameda, S.; Masuda, H.; Tasaka, M. Lateral root formation is blocked by a gain-of-function mutation in the SOLITARY-ROOT/IAA14 gene of Arabidopsis. Plant J.; 2002; 29, pp. 153-168. [DOI: https://dx.doi.org/10.1046/j.0960-7412.2001.01201.x] [PubMed: https://www.ncbi.nlm.nih.gov/pubmed/11862947]

251. Nagpal, P.; Walker, L.M.; Young, J.C.; Sonawala, A.; Timpte, C.; Estelle, M.; Reed, J.W. AXR2 encodes a member of the Aux/IAA protein family. Plant Physiol.; 2000; 123, pp. 563-574. [DOI: https://dx.doi.org/10.1104/pp.123.2.563] [PubMed: https://www.ncbi.nlm.nih.gov/pubmed/10859186]

252. Ouellet, F.; Overvoorde, P.J.; Theologis, A. IAA17/AXR3: Biochemical insight into an auxin mutant phenotype. Plant Cell; 2001; 13, pp. 829-841. [DOI: https://dx.doi.org/10.1105/tpc.13.4.829] [PubMed: https://www.ncbi.nlm.nih.gov/pubmed/11283339]

253. Yu, Z.; Zhang, F.; Friml, J.; Ding, Z. Auxin signaling: Research advances over the past 30 years. J. Integr. Plant Biol.; 2022; 64, pp. 371-392. [DOI: https://dx.doi.org/10.1111/jipb.13225] [PubMed: https://www.ncbi.nlm.nih.gov/pubmed/35018726]

254. Lavy, M.; Estelle, M. Mechanisms of auxin signaling. Development; 2016; 143, pp. 3226-3229. [DOI: https://dx.doi.org/10.1242/dev.131870]

255. Napier, R. The story of auxin-binding protein 1 (ABP1). Cold Spring Harb. Perspect. Biol.; 2021; 13, a039909. [DOI: https://dx.doi.org/10.1101/cshperspect.a039909] [PubMed: https://www.ncbi.nlm.nih.gov/pubmed/33903152]

256. Grones, P.; Chen, X.; Simon, S.; Kaufmann, W.A.; De Rycke, R.; Nodzyński, T.; Zažímalová, E.; Friml, J. Auxin-binding pocket of ABP1 is crucial for its gain-of-function cellular and developmental roles. J. Exp. Bot.; 2015; 66, pp. 5055-5065. [DOI: https://dx.doi.org/10.1093/jxb/erv177] [PubMed: https://www.ncbi.nlm.nih.gov/pubmed/25922490]

257. Michalko, J.; Dravecká, M.; Bollenbach, T.; Friml, J.; Ostergaard, L.; Napier, R.M.; Luschnig, C. Embryo-lethal phenotypes in early abp1 mutants are due to disruption of the neighboring BSM gene. F1000Research; 2015; 4, 1104. [DOI: https://dx.doi.org/10.12688/f1000research.7143.1] [PubMed: https://www.ncbi.nlm.nih.gov/pubmed/26629335]

258. Gao, Y.; Zhang, Y.; Zhang, D.; Dai, X.; Estelle, M.; Zhao, Y. Auxin binding protein 1 (ABP1) is not required for either auxin signaling or Arabidopsis development. Proc. Natl. Acad. Sci. USA; 2015; 112, pp. 2275-2280. [DOI: https://dx.doi.org/10.1073/pnas.1500365112] [PubMed: https://www.ncbi.nlm.nih.gov/pubmed/25646447]

259. Gelová, Z.; Gallei, M.; Pernisová, M.; Brunoud, G.; Zhang, X.; Glanc, M.; Li, L.; Michalko, J.; Pavlovičová, Z.; Verstraeten, I. et al. Developmental roles of auxin binding protein 1 in Arabidopsis thaliana. Plant Sci.; 2012; 303, 110750. [DOI: https://dx.doi.org/10.1016/j.plantsci.2020.110750]

260. Friml, J.; Gallei, M.; Gelová, Z.; Johnson, A.; Mazur, E.; Monzer, A.; Rodriquez, L.; Roosjen, M.; Verstraeten, I.; Živanovič, B.D. et al. ABP1–TMK auxin perception for global phosphorylation and auxin canalization. Nature; 2022; 609, pp. 575-581. [DOI: https://dx.doi.org/10.1038/s41586-022-05187-x]

261. Fiedler, L.; Friml, J. Rapid auxin signaling: Unknowns old and new. Curr. Opin. Plant Biol.; 2023; 75, 102443. [DOI: https://dx.doi.org/10.1016/j.pbi.2023.102443] [PubMed: https://www.ncbi.nlm.nih.gov/pubmed/37666097]

262. Timpte, C. Auxin binding protein: Curiouser and curiouser. Trends Plant Sci.; 2001; 6, pp. 586-590. [DOI: https://dx.doi.org/10.1016/S1360-1385(01)02150-1]

263. Napier, R.M.; David, K.M.; Perrot-Rechenmann, C. A short history of auxin-binding proteins. Plant Mol. Biol.; 2002; 49, pp. 339-348. [DOI: https://dx.doi.org/10.1023/A:1015259130955] [PubMed: https://www.ncbi.nlm.nih.gov/pubmed/12036258]

264. Badescu, G.O.; Napier, R.M. Receptors for auxin: Will it all end in TIRs?. Trends Plant Sci.; 2006; 11, pp. 217-223. [DOI: https://dx.doi.org/10.1016/j.tplants.2006.03.001] [PubMed: https://www.ncbi.nlm.nih.gov/pubmed/16564202]

265. Kim, S.H.; Bahk, S.; Nguyen, N.T.; Pham, M.L.A.; Kadam, U.S.; Hong, J.C.; Chung, W.S. Phosphorylation of the auxin signaling transcriptional repressor IAA15 by MPKs is required for the suppression of root development under drought stress in Arabidopsis. Nucleic Acids Res.; 2022; 50, pp. 10544-10561. [DOI: https://dx.doi.org/10.1093/nar/gkac798] [PubMed: https://www.ncbi.nlm.nih.gov/pubmed/36161329]

266. Kuhn, A.; Roosjen, M.; Mutte, S.; Dubey, S.M.; Carrillo Carrasco, V.P.; Boeren, S.; Monzer, A.; Koehorst, J.; Kohchi, T.; Nishihama, R. et al. RAF-like protein kinases mediate a deeply conserved, rapid auxin response. Cell; 2024; 187, pp. 130-148.e17. [DOI: https://dx.doi.org/10.1016/j.cell.2023.11.021]

267. Braun, N.; Wyrzykowska, J.; Muller, P.; David, K.; Couch, D.; Perrot-Rechenmann, C.; Fleming, A.J. Conditional repression of AUXIN BINDING PROTEIN1 reveals that it coordinates cell division and cell expansion during postembryonic shoot development in Arabidopsis and tobacco. Plant Cell; 2008; 10, pp. 2746-2762. [DOI: https://dx.doi.org/10.1105/tpc.108.059048]

268. Tromas, A.; Braun, N.; Muller, P.; Khodus, T.; Paponov, I.A.; Palme, K.; Ljung, K.; Lee, J.Y.; Benfey, P.; Murray, J.A. et al. The AUXIN BINDING PROTEIN 1 is required for differential auxin responses mediating root growth. PLoS ONE; 2009; 4, e6648. [DOI: https://dx.doi.org/10.1371/journal.pone.0006648]

269. Chen, J.G.; Ullah, H.; Young, J.C.; Sussman, M.R.; Jones, A.M. ABP1 is required for organized cell elongation and division in Arabidopsis embryogenesis. Genes Dev.; 2001; 15, pp. 902-911. [DOI: https://dx.doi.org/10.1101/gad.866201]

270. Emenecker, R.J.; Strader, L.C. Auxin-Abscisic Acid Interactions in Plant Growth and Development. Biomolecules; 2020; 10, 281. [DOI: https://dx.doi.org/10.3390/biom10020281] [PubMed: https://www.ncbi.nlm.nih.gov/pubmed/32059519]

271. Rück, A.; Pälme, K.; Venis, M.A.; Napier, R.M.; Felle, R.H. Patch-Clamp analysis establishes a role for an auxin binding protein in the auxin stimulation of plasma membrane current in Zea mays protoplasts. Plant J.; 1993; 4, pp. 41-46. [DOI: https://dx.doi.org/10.1046/j.1365-313X.1993.04010041.x]

272. Philippar, K.; Fuchs, I.; Luthen, H.; Hoth, S.; Bauer, C.S.; Haga, K.; Thiel, G.; Ljung, K.; Sandberg, G.; Bottger, M. et al. Auxin-induced K+ channel expression represents an essential step in coleoptile growth and gravitropism. Proc. Natl. Acad. Sci. USA; 1999; 96, pp. 12186-12191. [DOI: https://dx.doi.org/10.1073/pnas.96.21.12186]

273. Frias, I.; Caldeira, M.T.; Perez-Castineira, J.R.; Navarro-Avino, J.P.; Culianez-Macia, F.A.; Kuppinger, O.; Stransky, H.; Pages, M.; Hager, A.; Serrano, R. A major isoform of the maize plasma membrane H(+)-ATPase: Characterization and induction by auxin in coleoptiles. Plant Cell; 1996; 8, pp. 1533-1544. [DOI: https://dx.doi.org/10.1105/tpc.8.9.1533]

274. Spartz, A.K.; Ren, H.; Park, M.Y.; Grandt, K.N.; Lee, S.H.; Murphy, A.S.; Sussman, M.R.; Overvoorde, P.J.; Gray, W.M. SAUR Inhibition of PP2C-D Phosphatases Activates Plasma Membrane H+-ATPases to Promote Cell Expansion in Arabidopsis. Plant Cell; 2014; 26, pp. 2129-2142. [DOI: https://dx.doi.org/10.1105/tpc.114.126037]

275. Ren, H.; Gray, W.M. SAUR Proteins as Effectors of Hormonal and Environmental Signals in Plant Growth. Mol. Plant; 2015; 8, pp. 1153-1164. [DOI: https://dx.doi.org/10.1016/j.molp.2015.05.003] [PubMed: https://www.ncbi.nlm.nih.gov/pubmed/25983207]

276. Perrot-Rechenmann, C. Cellular responses to auxin: Division versus expansion. Cold Spring Harb. Perspect. Biol.; 2010; 2, a001446. [DOI: https://dx.doi.org/10.1101/cshperspect.a001446]

277. Liu, X.; Zhang, H.; Zhao, Y.; Feng, Z.; Li, Q.; Yang, H.Q.; Luan, S.; Li, J.; He, Z.H. Auxin controls seed dormancy through stimulation of abscisic acid signaling by inducing ARF-mediated ABI3 activation in Arabidopsis. Proc. Natl. Acad. Sci. USA; 2013; 110, pp. 15485-15490. [DOI: https://dx.doi.org/10.1073/pnas.1304651110] [PubMed: https://www.ncbi.nlm.nih.gov/pubmed/23986496]

278. Belin, C.; Megies, C.; Hauserova, E.; Lopez-Molina, L. Abscisic acid represses growth of the Arabidopsis embryonic axis after germination by enhancing auxin signaling. Plant Cell; 2009; 21, pp. 2253-2268. [DOI: https://dx.doi.org/10.1105/tpc.109.067702] [PubMed: https://www.ncbi.nlm.nih.gov/pubmed/19666738]

279. Thole, J.M.; Beisner, E.R.; Liu, J.; Venkova, S.V.; Strader, L.C. Abscisic acid regulates root elongation through the activities of auxin and ethylene in Arabidopsis. G3 Genes Genomes Genet.; 2014; 4, pp. 1259-1274. [DOI: https://dx.doi.org/10.1534/g3.114.011080] [PubMed: https://www.ncbi.nlm.nih.gov/pubmed/24836325]

280. Kazan, K. Auxin and the integration of environmental signals into plant root development. Ann. Bot.; 2013; 112, pp. 1655-1665. [DOI: https://dx.doi.org/10.1093/aob/mct229]

281. Jing, H.; Wilkinson, E.G.; Sageman-Furnas, K.; Strader, L.C. Auxin and abiotic stress responses. J. Exp. Bot.; 2023; 74, pp. 7000-7014. [DOI: https://dx.doi.org/10.1093/jxb/erad325] [PubMed: https://www.ncbi.nlm.nih.gov/pubmed/37591508]

282. Du, H.; Wu, N.; Chang, Y.; Li, X.; Xiao, J.; Xiong, L. Carotenoid deficiency impairs ABA and IAA biosynthesis and differentially affects drought and cold tolerance in rice. Plant Mol. Biol.; 2013; 83, pp. 475-488. [DOI: https://dx.doi.org/10.1007/s11103-013-0103-7]

283. Du, H.; Wu, N.; Fu, J.; Wang, S.; Li, X.; Xiao, J.; Xiong, L. A GH3 family member, OsGH3-2, modulates auxin and abscisic acid levels and differentially affects drought and cold tolerance in rice. J. Exp. Bot.; 2012; 63, pp. 6467-6480. [DOI: https://dx.doi.org/10.1093/jxb/ers300]

284. Kinoshita, N.; Wang, H.; Kasahara, H.; Liu, J.; Macpherson, C.; Machida, Y.; Kamiya, Y.; Hananh, M.A.; Chua, N.-H. IAA-Ala Resistant3, an evolutionarily conserved target of miR167, mediates Arabidopsis root architecture changes during high osmotic stress. Plant Cell; 2012; 24, pp. 3590-3602. [DOI: https://dx.doi.org/10.1105/tpc.112.097006] [PubMed: https://www.ncbi.nlm.nih.gov/pubmed/22960911]

285. Zhang, Y.; Li, Y.; Hassan, M.J.; Li, Z.; Peng, Y. Indole-3-acetic acid improves drought tolerance of white clover via activating auxin, abscisic acid and jasmonic acid related genes and inhibiting senescence genes. BMC Plant Biol.; 2020; 20, 150. [DOI: https://dx.doi.org/10.1186/s12870-020-02354-y] [PubMed: https://www.ncbi.nlm.nih.gov/pubmed/32268884]

286. Shi, H.; Chen, L.; Ye, T.; Liu, X.; Ding, K.; Chan, Z. Modulation of auxin content in Arabidopsis confers improved drought stress resistance. Plant Physiol. Biochem.; 2014; 82, pp. 209-217. [DOI: https://dx.doi.org/10.1016/j.plaphy.2014.06.008] [PubMed: https://www.ncbi.nlm.nih.gov/pubmed/24992887]

287. Sharma, A.; Gupta, A.; Ramakrishnan, M.; Ha, C.V.; Zheng, B.; Bhardwaj, M.; Tran, L.P. Roles of abscisic acid and auxin in plants during drought: A molecular point of view. Plant Physiol. Biochem.; 2023; 204, 108129. [DOI: https://dx.doi.org/10.1016/j.plaphy.2023.108129]

288. Balcerowicz, M.; Ranjan, A.; Rupprecht, L.; Fiene, G.; Hoecker, U. Auxin represses stomatal development in dark-grown seedlings via Aux/IAA proteins. Development; 2014; 141, pp. 3165-3176. [DOI: https://dx.doi.org/10.1242/dev.109181] [PubMed: https://www.ncbi.nlm.nih.gov/pubmed/25063454]

289. Le, J.; Liu, X.G.; Yang, K.Z.; Chen, X.L.; Zou, J.J.; Wang, H.Z.; Wang, M.; Vanneste, S.; Morita, M.; Tasaka, M. et al. Auxin transport and activity regulate stomatal patterning and development. Nat. Commun.; 2014; 5, 3090. [DOI: https://dx.doi.org/10.1038/ncomms4090] [PubMed: https://www.ncbi.nlm.nih.gov/pubmed/24463772]

290. Zhang, J.Y.; He, S.B.; Li, L.; Yang, H.Q. Auxin inhibits stomatal development through MONOPTEROS repression of a mobile peptide gene STOMAGEN in mesophyll. Proc. Natl. Acad. Sci. USA; 2014; 111, pp. E3015-E3023. [DOI: https://dx.doi.org/10.1073/pnas.1400542111] [PubMed: https://www.ncbi.nlm.nih.gov/pubmed/25002510]

291. Balcerowicz, M.; Hoecker, U. Auxin—A novel regulator of stomata differentiation. Trends Plant Sci.; 2014; 19, pp. 747-749. [DOI: https://dx.doi.org/10.1016/j.tplants.2014.10.006] [PubMed: https://www.ncbi.nlm.nih.gov/pubmed/25458848]

292. Kou, X.; Han, W.; Kang, J. Responses of root system architecture to water stress at multiple levels: A meta-analysis of trials under controlled conditions. Front. Plant Sci.; 2022; 13, 1085409. [DOI: https://dx.doi.org/10.3389/fpls.2022.1085409]

293. Mangano, S.; Denita-Juarez, S.P.; Choi, H.S.; Marzol, E.; Hwang, Y.; Ranocha, P.; Velasquez, S.M.; Borassi, C.; Barberini, M.L.; Aptekmann, A.A. et al. Molecular link between auxin and ROS-mediated polar growth. Proc. Natl. Acad. Sci. USA; 2017; 114, pp. 5289-5294. [DOI: https://dx.doi.org/10.1073/pnas.1701536114]

294. Löfke, C.; Zwiewka, M.; Heilmann, I.; Van Montagu, M.C.E.; Teichmann, T.; Friml, J. Asymmetric gibberellin signaling regulates vacuolar trafficking of PIN auxin transporters during root gravitropism. Proc. Natl. Acad. Sci. USA; 2013; 110, pp. 3627-3632. [DOI: https://dx.doi.org/10.1073/pnas.1300107110] [PubMed: https://www.ncbi.nlm.nih.gov/pubmed/23391733]

295. Choudhury, F.K.; Rivero, R.M.; Blumwald, E.; Mittler, R. Reactive oxygen species, abiotic stress and stress combination. Plant J.; 2017; 90, pp. 856-867. [DOI: https://dx.doi.org/10.1111/tpj.13299]

296. Saini, K.; AbdElgawad, H.; Markakis, M.N.; Schoenaers, S.; Asard, H.; Prinsen, E.; Beemster, G.T.S.; Vissenberg, K. Perturbation of Auxin homeostasis and signaling by PINOID overexpression induces stress responses in Arabidopsis. Front. Plant Sci.; 2017; 8, 1308. [DOI: https://dx.doi.org/10.3389/fpls.2017.01308] [PubMed: https://www.ncbi.nlm.nih.gov/pubmed/28824662]

297. Aloni, R. The induction of vascular tissues by auxin and cytokinin. Plant Hormones; Davies, P.J. Springer: Dordrecht, The Netherlands, 1995; pp. 531-546.

298. Werner, T.; Motyka, V.; Laucou, V.; Smets, R.; Onckelen, H.V.; Schmülling, T. Cytokinin-Defcient transgenic Arabidopsis plants show multiple developmental alterations indicating opposite functions of cytokinins in the regulation of shoot and root meristem activity. Plant Cell; 2003; 15, pp. 2532-2550. [DOI: https://dx.doi.org/10.1105/tpc.014928] [PubMed: https://www.ncbi.nlm.nih.gov/pubmed/14555694]

299. Hwang, I.; Sheen, J.; Müller, B. Cytokinin signaling networks. Annu. Rev. Plant Biol.; 2012; 63, pp. 353-380. [DOI: https://dx.doi.org/10.1146/annurev-arplant-042811-105503]

300. Cortleven, A.; Leuendorf, J.E.; Frank, M.; Pezzetta, D.; Bolt, S.; Schmulling, T. Cytokinin action in response to abiotic and biotic stresses in plants. Plant Cell Environ.; 2019; 42, pp. 998-1018. [DOI: https://dx.doi.org/10.1111/pce.13494] [PubMed: https://www.ncbi.nlm.nih.gov/pubmed/30488464]

301. Wang, W.; Hao, Q.; Wang, W.; Li, Q.; Chen, F.; Ni, W.Y.; Fu, D.; Wu, J.; Wang, W. The involvement of cytokinin and nitrogen metabolism in delayed fag leaf senescence in a wheat stay-green mutant, tasg1. Plant Sci.; 2019; 278, pp. 70-79. [DOI: https://dx.doi.org/10.1016/j.plantsci.2018.10.024] [PubMed: https://www.ncbi.nlm.nih.gov/pubmed/30471731]

302. Kurakawa, T.; Ueda, N.; Maekawa, M.; Kobayashi, K.; Kojima, M.; Nagato, Y.; Sakakibara, H.; Kyozuka, J. Direct control of shoot meristem activity by a cytokinin-activating enzyme. Nature; 2007; 445, pp. 652-655. [DOI: https://dx.doi.org/10.1038/nature05504]

303. Sang, Y.; Cheng, Z.; Zhang, X. Plant stem cells and de novo organogenesis. New Phytol.; 2018; 218, pp. 1334-1339. [DOI: https://dx.doi.org/10.1111/nph.15106] [PubMed: https://www.ncbi.nlm.nih.gov/pubmed/29574802]

304. Li, S.M.; Zheng, H.X.; Zhang, X.S.; Sui, N. Cytokinins as central regulators during plant growth and stress response. Plant Cell Rep.; 2021; 40, pp. 271-282. [DOI: https://dx.doi.org/10.1007/s00299-020-02612-1] [PubMed: https://www.ncbi.nlm.nih.gov/pubmed/33025178]

305. Gajdosová, S.; Spíchal, L.; Kamínek, M.; Hoyerová, K.; Novák, O.; Dobrev, P.I.; Galuszka, P.; Klíma, P.; Gaudinová, A.; Zizková, E. et al. Distribution, biological activities, metabolism, and the conceivable function of cis-zeatin-type cytokinins in plants. J. Exp. Bot.; 2011; 62, pp. 2827-2840. [DOI: https://dx.doi.org/10.1093/jxb/erq457] [PubMed: https://www.ncbi.nlm.nih.gov/pubmed/21282330]

306. Krall, L.; Raschke, M.; Zenk, M.H.; Baron, C. The Tzs protein from Agrobacterium tumefaciens C58 produces zeatin riboside 5′-phosphate from 4-hydroxy-3-methyl-2-(E)-butenyl diphos phate and AMP. FEBS Lett.; 2002; 527, pp. 315-318. [DOI: https://dx.doi.org/10.1016/S0014-5793(02)03258-1]

307. Werner, T.; Schmülling, T. Cytokinin action in plant development. Curr. Opin. Plant Biol.; 2009; 12, pp. 527-538. [DOI: https://dx.doi.org/10.1016/j.pbi.2009.07.002]

308. Hirose, N.; Takei, K.; Kuroha, T.; Kamada-Nobusada, T.; Hayashi, H.; Sakakibara, H. Regulation of cytokinin biosynthesis, compartmentalization and translocation. J. Exp. Bot.; 2008; 59, pp. 75-83. [DOI: https://dx.doi.org/10.1093/jxb/erm157]

309. Kamada-Nobusada, T.; Sakakibara, H. Molecular basis for cytokinin biosynthesis. Phytochemistry; 2009; 70, pp. 444-449. [DOI: https://dx.doi.org/10.1016/j.phytochem.2009.02.007] [PubMed: https://www.ncbi.nlm.nih.gov/pubmed/19285326]

310. Miyawaki, K.; Matsumoto-Kitano, M.; Kakimoto, T. Expression of cytokinin biosynthetic isopentenyltransferase genes in Arabidopsis: Tissue specificity and regulation by auxin, cytokinin, and nitrate. Plant J.; 2004; 37, pp. 128-138. [DOI: https://dx.doi.org/10.1046/j.1365-313X.2003.01945.x] [PubMed: https://www.ncbi.nlm.nih.gov/pubmed/14675438]

311. Duran-Medina, Y.; Diaz-Ramirez, D.; Marsch-Martinez, N. Cytokinins on the Move. Front. Plant Sci.; 2017; 8, 146. [DOI: https://dx.doi.org/10.3389/fpls.2017.00146] [PubMed: https://www.ncbi.nlm.nih.gov/pubmed/28228770]

312. Hwang, I.; Sheen, J. Two-component circuitry in Arabidopsis cytokinin signal transduction. Nature; 2001; 413, pp. 383-389. [DOI: https://dx.doi.org/10.1038/35096500]

313. Argueso, C.T.; Kieber, J.J. Cytokinin: From autoclaved DNA to two-component signaling. Plant Cell; 2024; 36, pp. 1429-1450. [DOI: https://dx.doi.org/10.1093/plcell/koad327] [PubMed: https://www.ncbi.nlm.nih.gov/pubmed/38163638]

314. Kakimoto, T. CKI1, a histidine kinase homolog implicated in cytokinin signal transduction. Science; 1996; 274, pp. 982-985. [DOI: https://dx.doi.org/10.1126/science.274.5289.982] [PubMed: https://www.ncbi.nlm.nih.gov/pubmed/8875940]

315. Inoue, T.; Higuchi, M.; Hashimoto, Y.; Seki, M.; Kobayashi, M.; Kato, T.; Tabata, S.; Shinozaki, K.; Kakimoto, T. Identification of CRE1 as a cytokinin receptor from Arabidopsis. Nature; 2001; 409, pp. 1060-1063. [DOI: https://dx.doi.org/10.1038/35059117] [PubMed: https://www.ncbi.nlm.nih.gov/pubmed/11234017]

316. Suzuki, T.; Miwa, K.; Ishikawa, K.; Yamada, H.; Aiba, H.; Mizuno, T. The Arabidopsis sensor His-kinase, AHk4, can respond to cytokinins. Plant Cell Physiol.; 2001; 42, pp. 107-113. [DOI: https://dx.doi.org/10.1093/pcp/pce037]

317. Ueguchi, C.; Sato, S.; Kato, T.; Tabata, S. The AHK4 gene involved in the cytokinin-signaling pathway as a direct receptor molecule in Arabidopsis thaliana. Plant Cell Physiol.; 2001; 42, pp. 751-755. [DOI: https://dx.doi.org/10.1093/pcp/pce094] [PubMed: https://www.ncbi.nlm.nih.gov/pubmed/11479382]

318. Yamada, H.; Suzuki, T.; Terada, K.; Takei, K.; Ishikawa, K.; Miwa, K.; Yamashino, T.; Mizuno, T. The Arabidopsis AHK4 histidine kinase is a cytokinin-binding receptor that transduces cytokinin signals across the membrane. Plant Cell Physiol.; 2001; 42, pp. 1017-1023. [DOI: https://dx.doi.org/10.1093/pcp/pce127] [PubMed: https://www.ncbi.nlm.nih.gov/pubmed/11577198]

319. Stolz, A.; Riefler, M.; Lomin, S.N.; Achazi, K.; Romanov, G.A.; Schmülling, T. The specificity of cytokinin signalling in Arabidopsis thaliana is mediated by differing ligand affinities and expression profiles of the receptors. Plant J.; 2011; 67, pp. 157-168. [DOI: https://dx.doi.org/10.1111/j.1365-313X.2011.04584.x]

320. Caesar, K.; Thamm, A.M.K.; Witthöft, J.; Elgass, K.; Huppenberger, P.; Grefen, C.; Horak, J.; Harter, K. Evidence for the localization of the Arabidopsis cytokinin receptors AHK3 and AHK4 in the endoplasmic reticulum. J. Exp. Bot.; 2011; 62, pp. 5571-5580. [DOI: https://dx.doi.org/10.1093/jxb/err238] [PubMed: https://www.ncbi.nlm.nih.gov/pubmed/21841169]

321. Lomin, S.N.; Yonekura-Sakakibara, K.; Romanov, G.A.; Sakakibara, H. Ligand-binding properties and subcellular localization of maize cytokinin receptors. J. Exp. Bot.; 2011; 62, pp. 5149-5159. [DOI: https://dx.doi.org/10.1093/jxb/err220] [PubMed: https://www.ncbi.nlm.nih.gov/pubmed/21778179]

322. Wulfetange, K.; Lomin, S.N.; Romanov, G.A.; Stolz, A.; Heyl, A.; Schmülling, T. The cytokinin receptors of Arabidopsis are located mainly to the endoplasmic reticulum. Plant Physiol.; 2011; 156, pp. 1808-1818. [DOI: https://dx.doi.org/10.1104/pp.111.180539]

323. Kubiasová, K.; Montesinos, J.C.; Šamajová, O.; Nisler, J.; Mik, V.; Semerádová, H.; Plíhalová, L.; Novák, O.; Marhavý, P.; Cavallari, N. et al. Cytokinin fluoroprobe reveals multiple sites of cytokinin perception at plasma membrane and endoplasmic reticulum. Nat Commun.; 2020; 11, 4285. [DOI: https://dx.doi.org/10.1038/s41467-020-17949-0] [PubMed: https://www.ncbi.nlm.nih.gov/pubmed/32855390]

324. Arkhipov, D.V.; Lomin, S.N.; Myakushina, Y.A.; Savelieva, E.M.; Osolodkin, D.I.; Romanov, G.A. Modeling of protein–protein interactions in cytokinin signal transduction. Int. J. Mol. Sci.; 2019; 20, pp. 2096-2118. [DOI: https://dx.doi.org/10.3390/ijms20092096] [PubMed: https://www.ncbi.nlm.nih.gov/pubmed/31035389]

325. Feng, J.; Wang, C.; Chen, Q.; Chen, H.; Ren, B.; Li, X.; Zuo, J. S-nitrosylation of phosphotransfer proteins represses cytokinin signaling. Nat. Commun.; 2013; 4, 1529. [DOI: https://dx.doi.org/10.1038/ncomms2541]

326. Miyata, S.; Urao, T.; Yamaguchi-Shinozaki, K.; Shinozaki, K. Characterization of genes for two-component phosphorelay mediators with a single HPt domain in Arabidopsis thaliana. FEBS Lett.; 1998; 437, pp. 11-14. [DOI: https://dx.doi.org/10.1016/S0014-5793(98)01188-0] [PubMed: https://www.ncbi.nlm.nih.gov/pubmed/9804162]

327. Schaller, G.E.; Kieber, J.J.; Shiu, S.-H. Two-component signaling elements and histidyl-aspartyl phosphorelays. Arab. Book Am. Soc. Plant Biol.; 2008; 6, e0112. [DOI: https://dx.doi.org/10.1199/tab.0112] [PubMed: https://www.ncbi.nlm.nih.gov/pubmed/22303237]

328. Suzuki, T.; Imamura, A.; Ueguchi, C.; Mizuno, T. Histidine-containing phosphotransfer (HPt) signal transducers Implicated in His-to-Asp phosphorelay in Arabidopsis. Plant Cell Physiol.; 1998; 39, pp. 1258-1268. [DOI: https://dx.doi.org/10.1093/oxfordjournals.pcp.a029329] [PubMed: https://www.ncbi.nlm.nih.gov/pubmed/10050311]

329. Suzuki, T.; Zakurai, K.; Imamura, A.; Nakamura, A.; Ueguchi, C.; Mizuno, T. Compilation and characterization of histidine-containing phosphotransmitters implicated in His-to-Asp phosphorelay in plants: AHP signal transducers of Arabidopsis thaliana. Biosci. Biotechnol. Biochem.; 2000; 64, pp. 2482-2485. [DOI: https://dx.doi.org/10.1271/bbb.64.2486] [PubMed: https://www.ncbi.nlm.nih.gov/pubmed/11193423]

330. Bishopp, A.; Help, H.; El-Showk, S.; Weijers, D.; Scheres, B.; Friml, J.; Benková, E.; Mähönen, A.; Helariutta, Y. A mutually inhibitory interaction between auxin and cytokinin specifies vascular pattern in roots. Curr. Biol.; 2011; 21, pp. 927-932. [DOI: https://dx.doi.org/10.1016/j.cub.2011.04.049]

331. Mähönen, A.P.; Bishopp, A.; Higuchi, M.; Nieminen, K.M.; Kinoshita, K.; Tormakangas, K.; Ikeda, Y.; Oka, A.; Kakimoto, T.; Helariutta, Y. Cytokinin signaling and its inhibitor AHP6 regulate cell fate during vascular development. Science; 2006; 311, pp. 94-98. [DOI: https://dx.doi.org/10.1126/science.1118875] [PubMed: https://www.ncbi.nlm.nih.gov/pubmed/16400151]

332. Besnard, F.; Refahi, Y.; Morin, V.; Marteaux, B.; Brunoud, G.; Chambrier, P.; Rozier, F.; Mirabet, V.; Legrand, J.; Lainé, S. et al. Cytokinin signalling inhibitory fields provide robustness to phyllotaxis. Nature; 2014; 505, pp. 417-421. [DOI: https://dx.doi.org/10.1038/nature12791]

333. Moreira, S.; Bishopp, A.; Carvalho, H.; Campilho, A. AHP6 inhibits cytokinin signaling to regulate the orientation of pericycle cell division during lateral root initiation. PLoS ONE; 2013; 8, e56370. [DOI: https://dx.doi.org/10.1371/journal.pone.0056370]

334. To, J.P.C.; Kieber, J.J. Cytokinin signaling: Two-components and more. Trends Plant Sci.; 2008; 13, pp. 85-92. [DOI: https://dx.doi.org/10.1016/j.tplants.2007.11.005] [PubMed: https://www.ncbi.nlm.nih.gov/pubmed/18262459]

335. Argyros, R.D.; Mathews, D.E.; Chiang, Y.-H.; Palmer, C.M.; Thibault, D.M.; Etheridge, N.; Argyros, D.A.; Mason, M.G.; Kieber, J.J.; Schaller, G.E. Type B response regulators of Arabidopsis play key roles in cytokinin signaling and plant development. Plant Cell; 2008; 20, pp. 2102-2116. [DOI: https://dx.doi.org/10.1105/tpc.108.059584]

336. Ishida, K.; Yamashino, T.; Yokoyama, A.; Mizuno, T. Three Type-B Response Regulators, ARR1, ARR10 and ARR12, play essential but redundant roles in cytokinin signal transduction throughout the life cycle of Arabidopsis thaliana. Plant Cell Physiol.; 2008; 49, pp. 47-57. [DOI: https://dx.doi.org/10.1093/pcp/pcm165] [PubMed: https://www.ncbi.nlm.nih.gov/pubmed/18037673]

337. Kim, H.J.; Kieber, J.J.; Schaller, G.E. The rice F-box protein KISS ME DEADLY2 functions as a negative regulator of cytokinin signalling. Plant Signal. Behav.; 2013; 8, e26434. [DOI: https://dx.doi.org/10.4161/psb.26434]

338. Kim, K.; Ryu, H.; Cho, Y.-H.; Scacchi, E.; Sabatini, S.; Hwang, I. Cytokinin-facilitated proteolysis of ARABIDOPSIS RESPONSE REGULATOR 2 attenuates signaling output in two-component circuitry. Plant J.; 2012; 69, pp. 934-945. [DOI: https://dx.doi.org/10.1111/j.1365-313X.2011.04843.x]

339. Brandstatter, I.; Kieber, J.J. Two genes with similarity to bacterial response regulators are rapidly and specifically induced by cytokinin in Arabidopsis. Plant Cell; 1998; 10, pp. 1009-1020. [DOI: https://dx.doi.org/10.1105/tpc.10.6.1009] [PubMed: https://www.ncbi.nlm.nih.gov/pubmed/9634588]

340. D’Agostino, I.B.; Deruère, J.; Kieber, J.J. Characterization of the response of the Arabidopsis ARR gene family to cytokinin. Plant Physiol.; 2000; 124, pp. 1706-1717. [DOI: https://dx.doi.org/10.1104/pp.124.4.1706]

341. Taniguchi, M.; Kiba, T.; Sakakibara, H.; Ueguchi, C.; Mizuno, T.; Sugiyama, T. Expression of Arabidopsis response regulator homologs is induced by cytokinins and nitrate. FEBS Lett.; 1998; 429, pp. 259-262. [DOI: https://dx.doi.org/10.1016/S0014-5793(98)00611-5] [PubMed: https://www.ncbi.nlm.nih.gov/pubmed/9662428]

342. Taniguchi, M.; Sasaki, N.; Tsuge, T.; Aoyama, T.; Oka, A. ARR1 directly activates cytokinin response genes that encode proteins with diverse regulatory functions. Plant Cell Physiol.; 2007; 48, pp. 263-277. [DOI: https://dx.doi.org/10.1093/pcp/pcl063] [PubMed: https://www.ncbi.nlm.nih.gov/pubmed/17202182]

343. Meng, W.J.; Cheng, Z.J.; Sang, Y.L.; Zhang, M.M.; Rong, X.F.; Wang, Z.W.; Tang, Y.Y.; Zhang, X.S. Type-B ARABIDOPSIS RESPONSE REGULATORs specify the shoot stem cell niche by dual regulation of WUSCHEL. Plant Cell; 2017; 29, pp. 1357-1372. [DOI: https://dx.doi.org/10.1105/tpc.16.00640] [PubMed: https://www.ncbi.nlm.nih.gov/pubmed/28576846]

344. Wang, J.; Tian, C.; Zhang, C.; Shi, B.; Cao, X.; Zhang, T.-Q.; Zhao, Z.; Wang, J.-W.; Jiao, Y. Cytokinin signaling activates WUSCHEL expression during axillary meristem initiation. Plant Cell; 2017; 29, pp. 1373-1387. [DOI: https://dx.doi.org/10.1105/tpc.16.00579] [PubMed: https://www.ncbi.nlm.nih.gov/pubmed/28576845]

345. Zhang, F.; May, A.; Irish, V.F. Type-B ARABIDOPSIS RESPONSE REGULATORs Directly Activate WUSCHEL. Trends Plant Sci.; 2017; 22, pp. 815-817. [DOI: https://dx.doi.org/10.1016/j.tplants.2017.08.007]

346. Zubo, Y.; Blakley, I.C.; Yamburenko, M.; Worthen, J.M.; Street, I.; Franco-Zorrilla, J.M.; Zhang, W.; Hill, K.; Raines, T.; Solano, R. et al. Cytokinin induces genome-wide binding of the type-B response regulator ARR10 to regulate growth and development in Arabidopsis. Proc. Natl. Acad. Sci. USA; 2017; 114, pp. E5995-E6004. [DOI: https://dx.doi.org/10.1073/pnas.1620749114]

347. Zwack, P.J.; De Clercq, I.; Howton, T.C.; Hallmark, H.T.; Hurny, A.; Keshishian, E.A.; Parish, A.M.; Benkova, E.; Mukhtar, M.S.; Van Breusegem, F. et al. Cytokinin Response Factor 6 Represses Cytokinin-Associated Genes during Oxidative Stress. Plant Physiol.; 2016; 172, pp. 1249-1258. [DOI: https://dx.doi.org/10.1104/pp.16.00415] [PubMed: https://www.ncbi.nlm.nih.gov/pubmed/27550996]

348. Keshishian, E.A.; Rashotte, A.M. Plant cytokinin signalling. Essays Biochem.; 2015; 58, pp. 13-27. [DOI: https://dx.doi.org/10.1042/bse0580013]

349. Hallmark, H.T.; Rashotte, A.M. Review—Cytokinin response factors: Responding to more than cytokinin. Plant Sci.; 2019; 289, 110251. [DOI: https://dx.doi.org/10.1016/j.plantsci.2019.110251]

350. Gentile, D.; Serino, G.; Frugis, G. CRF transcription factors in the trade-off between abiotic stress response and plant developmental processes. Front. Genet.; 2024; 15, 1377204. [DOI: https://dx.doi.org/10.3389/fgene.2024.1377204] [PubMed: https://www.ncbi.nlm.nih.gov/pubmed/38694876]

351. Hasanuzzaman, M.; Bhuyan, M.H.M.B.; Zulfiqar, F.; Raza, A.; Mohsin, S.M.; Mahmud, J.A.; Fujita, M.; Fotopoulos, V. Reactive oxygen species and antioxidant defense in plants under abiotic stress: Revisiting the crucial role of a universal defense regulator. Antioxidants; 2020; 9, 681. [DOI: https://dx.doi.org/10.3390/antiox9080681]

352. Maurya, A.K. Oxidative stress in crop plants. Agronomic Crops: Volume 3: Stress Responses and Tolerance; Hasanuzzaman, M. Springer: Singapore, 2020; Volume 3, pp. 349-380. [DOI: https://dx.doi.org/10.1007/978-981-15-0025-1_18]

353. Šimášková, M.; O’Brien, J.A.; Khan, M.; Van Noorden, G.; Ötvös, K.; Vieten, A.; De Clercq, I.; Van Haperen, J.M.A.; Cuesta, C.; Hoyerová, K. et al. Cytokinin response factors regulate PIN-FORMED auxin transporters. Nat. Commun.; 2015; 6, 8717. [DOI: https://dx.doi.org/10.1038/ncomms9717]

354. DelloIoio, R.; Nakamura, K.; Moubayidin, L.; Perilli, S.; Taniguchi, M.; Morita, M.T.; Aoyama, T.; Costantino, P.; Sabatini, S. A genetic framework for the control of cell division and differentiation in the root meristem. Science; 2008; 322, pp. 1380-1384. [DOI: https://dx.doi.org/10.1126/science.1164147]

355. Pernisová, M.; Klíma, P.; Horák, J.; Válková, M.; Malbeck, J.; Soucek, P.; Reichman, P.; Hoyerová, K.; Dubová, J.; Friml, J. et al. Cytokinins modulate auxin-induced organogenesis in plants via regulation of the auxin efflux. Proc. Natl. Acad. Sci. USA; 2009; 106, pp. 3609-3614. [DOI: https://dx.doi.org/10.1073/pnas.0811539106] [PubMed: https://www.ncbi.nlm.nih.gov/pubmed/19211794]

356. Ruzicka, K.; Simásková, M.; Duclercq, J.; Petrásek, J.; Zazímalová, E.; Simon, S.; Friml, J.; Van Montagu, M.C.; Benková, E. Cytokinin regulates root meristem activity via modulation of the polar auxin transport. Proc. Natl. Acad. Sci. USA; 2009; 106, pp. 4284-4289. [DOI: https://dx.doi.org/10.1073/pnas.0900060106] [PubMed: https://www.ncbi.nlm.nih.gov/pubmed/19246387]

357. Marhavý, P.; Bielach, A.; Abas, L.; Abuzeineh, A.; Duclercq, J.; Tanaka, H.; Pařezová, M.; Petrášek, J.; Friml, J.; Kleine-Vehn, J. et al. Cytokinin modulates endocytic trafficking of PIN1 auxin efflux carrier to control plant organogenesis. Dev. Cell; 2011; 21, pp. 796-804. [DOI: https://dx.doi.org/10.1016/j.devcel.2011.08.014]

358. Schlereth, A.; Möller, B.; Liu, W.; Kientz, M.; Flipse, J.; Rademacher, E.H.; Schmid, M.; Jürgens, G.; Weijers, D. MONOPTEROS controls embryonic root initiation by regulating a mobile transcription factor. Nature; 2010; 464, pp. 913-916. [DOI: https://dx.doi.org/10.1038/nature08836] [PubMed: https://www.ncbi.nlm.nih.gov/pubmed/20220754]

359. Ashikari, M.; Sakakibara, H.; Lin, S.; Yamamoto, T.; Takashi, T.; Nishimura, A.; Angeles, E.R.; Qian, Q.; Kitano, H.; Matsuoka, M. Cytokinin oxidase regulates rice grain production. Science; 2005; 309, pp. 741-745. [DOI: https://dx.doi.org/10.1126/science.1113373] [PubMed: https://www.ncbi.nlm.nih.gov/pubmed/15976269]

360. Bartrina, I.; Otto, E.; Strnad, M.; Werner, T.; Schmülling, T. Cytokinin regulates the activity of reproductive meristems, flower organ size, ovule formation, and thus seed yield in Arabidopsis thaliana. Plant Cell; 2011; 23, pp. 69-80. [DOI: https://dx.doi.org/10.1105/tpc.110.079079]

361. Cucinotta, M.; Manrique, S.; Guazzotti, A.; Quadrelli, N.E.; Mendes, M.A.; Benkova, E.; Colombo, L. Cytokinin response factors integrate auxin and cytokinin pathways for female reproductive organ development. Development; 2016; 143, pp. 4419-4424. [DOI: https://dx.doi.org/10.1242/dev.143545]

362. Marsch-Martínez, N.; Ramos-Cruz, D.; Reyes-Olalde, J.I.; Lozano-Sotomayor, P.; Zúñiga-Mayo, V.M.; de Folter, S. The role of cytokinin during Arabidopsis gynoecia and fruit morphogenesis and patterning. Plant J.; 2012; 72, pp. 222-234. [DOI: https://dx.doi.org/10.1111/j.1365-313X.2012.05062.x] [PubMed: https://www.ncbi.nlm.nih.gov/pubmed/22640521]

363. Müller, C.J.; Larsson, E.; Spíchal, L.; Sundberg, E. Cytokinin-auxin crosstalk in the gynoecial primordium ensures correct domain patterning. Plant Physiol.; 2017; 175, pp. 1144-1157. [DOI: https://dx.doi.org/10.1104/pp.17.00805]

364. Sehra, B.; Franks, R.G. Auxin and cytokinin act during gynoecial patterning and the development of ovules from the meristematic medial domain. Wiley Interdiscip. Rev. Dev. Biol.; 2015; 4, pp. 555-571. [DOI: https://dx.doi.org/10.1002/wdev.193] [PubMed: https://www.ncbi.nlm.nih.gov/pubmed/25951007]

365. Cheng, C.-Y.; Mathews, D.E.; Schaller, G.E.; Kieber, J.J. Cytokinin-dependent specification of the functional megaspore in the Arabidopsis female gametophyte. Plant J.; 2013; 73, pp. 929-940. [DOI: https://dx.doi.org/10.1111/tpj.12084]

366. Yuan, L.; Liu, Z.; Song, X.; Johnson, C.; Yu, X.; Sundaresan, V. The CKI1 histidine kinase specifies the female gametic precursor of the endosperm. Dev. Cell; 2016; 37, pp. 34-46. [DOI: https://dx.doi.org/10.1016/j.devcel.2016.03.009] [PubMed: https://www.ncbi.nlm.nih.gov/pubmed/27046830]

367. Immanen, J.; Nieminen, K.; Smolander, O.-P.; Kojima, M.; Alonso Serra, J.; Koskinen, P.; Zhang, J.; Elo, A.; Mähönen, A.P.; Street, N. et al. Cytokinin and auxin display distinct but interconnected distribution and signaling profiles to stimulate cambial activity. Curr. Biol.; 2016; 26, pp. 1990-1997. [DOI: https://dx.doi.org/10.1016/j.cub.2016.05.053] [PubMed: https://www.ncbi.nlm.nih.gov/pubmed/27426519]

368. Nieminen, K.; Immanen, J.; Laxell, M.; Kauppinen, L.; Tarkowski, P.; Dolezal, K.; Tähtiharju, S.; Elo, A.; Decourteix, M.; Ljung, K. et al. Cytokinin signaling regulates cambial development in poplar. Proc. Natl. Acad. Sci. USA; 2008; 105, pp. 20032-20037. [DOI: https://dx.doi.org/10.1073/pnas.0805617106] [PubMed: https://www.ncbi.nlm.nih.gov/pubmed/19064928]

369. Dello Ioio, R.; Galinha, C.; Fletcher, A.G.; Grigg, S.P.; Molnar, A.; Willemsen, V.; Scheres, B.; Sabatini, S.; Baulcombe, D.; Maini, P.K. et al. A PHABULOSA/cytokinin feedback loop controls root growth in Arabidopsis. Curr. Biol.; 2012; 22, pp. 1699-1704. [DOI: https://dx.doi.org/10.1016/j.cub.2012.07.005]

370. Moubayidin, L.; Perilli, S.; DelloIoio, R.; Di Mambro, R.; Costantino, P.; Sabatini, S. The rate of cell differentiation controls the Arabidopsis root meristem growth phase. Curr. Biol.; 2010; 20, pp. 1138-1143. [DOI: https://dx.doi.org/10.1016/j.cub.2010.05.035] [PubMed: https://www.ncbi.nlm.nih.gov/pubmed/20605455]

371. Werner, T.; Motyka, V.; Strnad, M.; Schmülling, T. Regulation of plant growth by cytokinin. Proc. Natl. Acad. Sci. USA; 2001; 98, pp. 10487-10492. [DOI: https://dx.doi.org/10.1073/pnas.171304098] [PubMed: https://www.ncbi.nlm.nih.gov/pubmed/11504909]

372. Werner, T.; Nehnevajova, E.; Köllmer, I.; Novák, O.; Strnad, M.; Krämer, U.; Schmülling, T. Root-specifc reduction of cytokinin causes enhanced root growth, drought tolerance, and leaf mineral enrichment in Arabidopsis and tobacco. Plant Cell; 2010; 22, pp. 3905-3920. [DOI: https://dx.doi.org/10.1105/tpc.109.072694]

373. Chang, L.; Ramireddy, E.; Schmülling, T. Lateral root formation and growth of Arabidopsis is redundantly regulated by cytokinin metabolism and signalling genes. J. Exp. Bot.; 2013; 64, pp. 5021-5032. [DOI: https://dx.doi.org/10.1093/jxb/ert291]

374. Chang, L.; Ramireddy, E.; Schmülling, T. Cytokinin as a positional cue regulating lateral root spacing in Arabidopsis. J. Exp. Bot.; 2015; 66, pp. 4759-4768. [DOI: https://dx.doi.org/10.1093/jxb/erv252]

375. Dello Ioio, R.D.; Linhares, F.S.; Scacchi, E.; Casamitjana-martinez, E.; Heidstra, R.; Costantino, P.; Sabatini, S. Cytokinins determine arabidopsis root-meristem size by controlling cell diferentiation. Curr. Biol.; 2007; 17, pp. 678-682. [DOI: https://dx.doi.org/10.1016/j.cub.2007.02.047]

376. Ren, B.; Liang, Y.; Deng, Y.; Chen, Q.; Zhang, J.; Yang, X.; Zuo, J. Genome-wide comparative analysis of type-A Arabidopsis response regulator genes by overexpression studies reveals their diverse roles and regulatory mechanisms in cytokinin signaling. Cell Res.; 2009; 19, pp. 1178-1190. [DOI: https://dx.doi.org/10.1038/cr.2009.88] [PubMed: https://www.ncbi.nlm.nih.gov/pubmed/19621034]

377. Sakamoto, T.; Sakakibara, H.; Kojima, M.; Yamamoto, Y.; Nagasaki, H.; Inukai, Y.; Sato, Y.; Matsuoka, M. Ectopic expression of KNOT TED1-like homeobox protein induces expression of cytokinin biosynthesis genes in rice. Plant Physiol.; 2006; 142, pp. 54-62. [DOI: https://dx.doi.org/10.1104/pp.106.085811] [PubMed: https://www.ncbi.nlm.nih.gov/pubmed/16861569]

378. Cheng, X.; Jiang, H.; Zhang, J.; Qian, Y.; Zhu, S.; Cheng, B. Over expression of type-A rice response regulators, OsRR3 and OsRR5, results in lower sensitivity to cytokinins. Genet. Mol. Res.; 2010; 9, pp. 348-359. [DOI: https://dx.doi.org/10.4238/vol9-1gmr739] [PubMed: https://www.ncbi.nlm.nih.gov/pubmed/20309821]

379. Choi, J.; Huh, S.U.; Kojima, M.; Sakakibara, H.; Paek, K.H.; Hwang, I. The Cytokinin-Activated Transcription Factor ARR2 Promotes Plant Immunity via TGA3/NPR1-Dependent Salicylic Acid Signaling in Arabidopsis. Dev. Cell; 2010; 19, pp. 284-295. [DOI: https://dx.doi.org/10.1016/j.devcel.2010.07.011]

380. Argueso, C.T.; Ferreira, F.J.; Epple, P.; To, J.P.; Hutchison, C.E.; Schaller, G.E.; Dangl, J.L.; Kieber, J.J. Two-Component Elements Mediate Interactions Between Cytokinin and Salicylic Acid in Plant Immunity. PLoS Genet.; 2012; 8, e1002448. [DOI: https://dx.doi.org/10.1371/journal.pgen.1002448] [PubMed: https://www.ncbi.nlm.nih.gov/pubmed/22291601]

381. Liang, Y.S.; Ermawati, N.; Cha, J.Y.; Jung, M.H.; Su’udi, H.; Kim, M.G.; Ha, S.H.; Park, C.G.; Son, D. Overexpression of an AP2/ERF-Type Transcription Factor CRF5 Confers Pathogen Resistance to Arabidopsis Plants. J. Korean Soc. Appl. Biol. Chem.; 2010; 53, pp. 142-148. [DOI: https://dx.doi.org/10.3839/jksabc.2010.024]

382. Marín-de la Rosa, N.; Pfeiffer, A.; Hill, K.; Locascio, A.; Bhalerao, R.P.; Miskolczi, P.; Grønlund, A.L.; Wanchoo-Kohli, A.; Thomas, S.G.; Bennett, M.J. et al. Genome Wide Binding Site Analysis Reveals Transcriptional Coactivation of Cytokinin-Responsive Genes by DELLA Proteins. PLoS Genet.; 2015; 11, e1005337. [DOI: https://dx.doi.org/10.1371/journal.pgen.1005337]

383. Chen, B.; Yang, H. 6-Benzylaminopurine alleviates chilling injury of postharvest cucumber fruit through modulating antioxidant system and energy status. J. Sci. Food Agric.; 2013; 93, pp. 1915-1921. [DOI: https://dx.doi.org/10.1002/jsfa.5990]

384. Rivero, R.M.; Kojima, M.; Gepstein, A.; Sakakibara, H.; Mittler, R.; Gepstein, S.; Blumwald, E. Delayed leaf senescence induces extreme drought tolerance in a flowering plant. Proc. Natl. Acad. Sci. USA; 2007; 104, pp. 19631-19636. [DOI: https://dx.doi.org/10.1073/pnas.0709453104]

385. Kamal, N.M.; Gorafi, Y.S.A.; Abdelrahman, M.; Abdellatef, E.; Tsujimoto, H. Stay-green trait: A prospective approach for yield potential, and drought and heat stress adaptation in globally important cereals. Int. J. Mol. Sci.; 2019; 20, 5837. [DOI: https://dx.doi.org/10.3390/ijms20235837]

386. Baldoni, E.; Frugis, G.; Martinelli, F.; Benny, J.; Paffetti, D.; Buti, M. A comparative transcriptomic meta-analysis revealed conserved key genes and regulatory networks involved in drought tolerance in cereal crops. Int. J. Mol. Sci.; 2021; 22, 13062. [DOI: https://dx.doi.org/10.3390/ijms222313062]

387. Tan, S.; Sha, Y.; Sun, L.; Li, Z. Abiotic stress-induced leaf senescence: Regulatory mechanisms and application. Int. J. Mol. Sci.; 2023; 24, 11996. [DOI: https://dx.doi.org/10.3390/ijms241511996]

388. Zwack, P.J.; Robinson, B.R.; Risley, M.G.; Rashotte, A.M. Cytokinin response factor 6 negatively regulates leaf senescence and is induced in response to cytokinin and numerous abiotic stresses. Plant Cell Physiol.; 2013; 54, pp. 971-981. [DOI: https://dx.doi.org/10.1093/pcp/pct049]

389. Swinka, C.; Hellmann, E.; Zwack, P.; Banda, R.; Rashotte, A.M.; Heyl, A. Cytokinin response factor 9 represses cytokinin responses in flower development. Int. J. Mol. Sci.; 2023; 24, 4380. [DOI: https://dx.doi.org/10.3390/ijms24054380] [PubMed: https://www.ncbi.nlm.nih.gov/pubmed/36901811]

390. Raines, T.; Shanks, C.; Cheng, C.-Y.; McPherson, D.; Argueso, C.T.; Kim, H.J.; Franco-Zorrilla, J.M.; López-Vidriero, I.; Solano, R.; Vaňková, R. et al. The cytokinin response factors modulate root and shoot growth and promote leaf senescence in Arabidopsis. Plant J.; 2016; 85, pp. 134-147. [DOI: https://dx.doi.org/10.1111/tpj.13097]

391. Kwon, T. Cytokinin Response Factor 2 positively regulates salicylic acid mediated plant immunity in Arabidopsis thaliana. Plant Biotechnol.; 2016; 33, pp. 207-210. [DOI: https://dx.doi.org/10.5511/plantbiotechnology.16.0711a]

392. Okazaki, K.; Kabeya, Y.; Suzuki, K.; Mori, T.; Ichikawa, T.; Matsui, M.; Nakanishi, H.; Miyagishima, S.Y. The PLASTID DIVISION1 and 2 components of the chloroplast division machinery determine the rate of chloroplast division in land plant cell differentiation. Plant Cell; 2009; 21, pp. 1769-1780. [DOI: https://dx.doi.org/10.1105/tpc.109.067785]

393. Vercruyssen, L.; Tognetti, V.B.; Gonzalez, N.; Van Dingenen, J.; De Milde, L.; Bielach, A.; De Rycke, R.; Van Breusegem, F.; Inzé, D. GROWTH REGULATING FACTOR5 stimulates Arabidopsis chloroplast division, photosynthesis, and leaf longevity. Plant Physiol.; 2015; 167, pp. 817-832. [DOI: https://dx.doi.org/10.1104/pp.114.256180] [PubMed: https://www.ncbi.nlm.nih.gov/pubmed/25604530]

394. Nishiyama, R.; Watanabe, Y.; Fujita, Y.; Le, D.T.; Kojima, M.; Werner, T.; Vankova, R.; Yamaguchi-Shinozaki, K.; Shinozaki, K.; Kakimoto, T. et al. Analysis of Cytokinin Mutants and Regulation of Cytokinin Metabolic Genes Reveals Important Regulatory Roles of Cytokinins in Drought, Salt and Abscisic Acid Responses, and Abscisic Acid Biosynthesis. Plant Cell; 2011; 23, pp. 2169-2183. [DOI: https://dx.doi.org/10.1105/tpc.111.087395] [PubMed: https://www.ncbi.nlm.nih.gov/pubmed/21719693]

395. Kang, N.Y.; Cho, C.; Kim, N.Y.; Kim, J. Cytokinin Receptor-Dependent and Receptor-Independent Pathways in the Dehydration Response of Arabidopsis thaliana. J. Plant Physiol.; 2012; 169, pp. 1382-1391. [DOI: https://dx.doi.org/10.1016/j.jplph.2012.05.007]

396. Nishiyama, R.; Watanabe, Y.; Leyva-Gonzalez, M.A.; Ha, C.V.; Fujita, Y.; Tanaka, M.; Seki, M.; Yamaguchi-Shinozaki, K.; Shinozaki, K.; Herrera-Estrella, L. et al. Arabidopsis AHP2, AHP3, and AHP5 Histidine Phosphotransfer Proteins Function as Redundant Negative Regulators of Drought Stress Response. Proc. Natl. Acad. Sci. USA; 2013; 110, pp. 4840-4845. [DOI: https://dx.doi.org/10.1073/pnas.1302265110] [PubMed: https://www.ncbi.nlm.nih.gov/pubmed/23487796]

397. Nguyen, K.H.; Ha, C.V.; Nishiyama, R.; Watanabe, Y.; Leyva-González, M.A.; Fujita, Y.; Tran, U.T.; Li, W.; Tanaka, M.; Seki, M. et al. Arabidopsis Type B Cytokinin Response Regulators ARR1, ARR10, and ARR12 Negatively Regulate Plant Responses to Drought. Proc. Natl. Acad. Sci. USA; 2016; 113, pp. 3090-3095. [DOI: https://dx.doi.org/10.1073/pnas.1600399113]

398. Mushtaq, N.; Wang, Y.; Fan, J.; Li, Y.; Ding, J. Down-Regulation of Cytokinin Receptor Gene SlHK2 Improves Plant Tolerance to Drought, Heat, and Combined Stresses in Tomato. Plants; 2022; 11, 154. [DOI: https://dx.doi.org/10.3390/plants11020154] [PubMed: https://www.ncbi.nlm.nih.gov/pubmed/35050042]

399. Melotto, M.; Underwood, W.; Koczan, J.; Nomura, K.; He, S.Y. Plant Stomata Function in Innate Immunity Against Bacterial Invasion. Cell; 2006; 126, pp. 969-980. [DOI: https://dx.doi.org/10.1016/j.cell.2006.06.054] [PubMed: https://www.ncbi.nlm.nih.gov/pubmed/16959575]

400. Guzel Deger, A.; Scherzer, S.; Nuhkat, M.; Kedzierska, J.; Kollist, H.; Brosché, M.; Unyayar, S.; Boudsocq, M.; Hedrich, R.; Roelfsema, M.R. Guard Cell SLAC1-Type Anion Channels Mediate Flagellin-Induced Stomatal Closure. New Phytol.; 2015; 208, pp. 162-173. [DOI: https://dx.doi.org/10.1111/nph.13435]

401. Nolan, T.M.; Vukašinović, N.; Liu, D.; Russinova, E.; Yin, Y. Brassinosteroids: Multidimensional Regulators of Plant Growth, Development, and Stress Responses. Plant Cell; 2020; 32, pp. 295-318. [DOI: https://dx.doi.org/10.1105/tpc.19.00335]

402. Grove, M.D.; Spencer, G.F.; Rohwedder, W.K.; Mandava, N.; Worley, J.F.; Warthen, J.D.; Steffens, G.L.; Flippen-Anderson, J.L.; Cook, J.C. Brassinolide, a plant growth-promoting steroid isolated from Brassica napus pollen. Nature; 1979; 281, pp. 216-217. [DOI: https://dx.doi.org/10.1038/281216a0]

403. Manghwar, H.; Hussain, A.; Ali, Q.; Liu, F. Brassinosteroids (BRs) Role in Plant Development and Coping with Different Stresses. Int. J. Mol. Sci.; 2022; 23, 1012. [DOI: https://dx.doi.org/10.3390/ijms23031012] [PubMed: https://www.ncbi.nlm.nih.gov/pubmed/35162936]

404. Li, J.; Nagpal, P.; Vitart, V.; McMorris, T.C.; Chory, J. A role for brassinosteroids in light-dependent development of Arabidopsis. Science; 1996; 272, pp. 398-401. [DOI: https://dx.doi.org/10.1126/science.272.5260.398] [PubMed: https://www.ncbi.nlm.nih.gov/pubmed/8602526]

405. Szekeres, M.; Németh, K.; Koncz-Kálmán, Z.; Mathur, J.; Kauschmann, A.; Altmann, T.; Rédei, G.P.; Nagy, F.; Schell, J.; Koncz, C. Brassinosteroids rescue the deficiency of CYP90, a cytochrome P450, controlling cell elongation and de-etiolation in Arabidopsis. Cell; 1996; 85, pp. 171-182. [DOI: https://dx.doi.org/10.1016/S0092-8674(00)81094-6]

406. Ren, H.; Willige, B.C.; Jaillais, Y.; Geng, S.; Park, M.Y.; Gray, W.M.; Chory, J. BRASSINOSTEROID-SIGNALING KINASE 3, a plasma membrane-associated scaffold protein involved in early brassinosteroid signaling. PLoS Genet.; 2019; 15, e1007904. [DOI: https://dx.doi.org/10.1371/journal.pgen.1007904] [PubMed: https://www.ncbi.nlm.nih.gov/pubmed/30615605]

407. Ying, W.; Wang, Y.; Wei, H.; Luo, Y.; Ma, Q.; Zhu, H.; Janssens, H.; Vukašinović, N.; Kvasnica, M.; Winne, J.M. et al. Structure and function of the Arabidopsis ABC transporter ABCB19 in brassinosteroid export. Science; 2024; 383, eadj4591. [DOI: https://dx.doi.org/10.1126/science.adj4591] [PubMed: https://www.ncbi.nlm.nih.gov/pubmed/38513023]

408. Clouse, S.; Langford, M.; McMorris, T.C. A brassinosteroid insensitive mutant in Arabidopsis thaliana exhibits multiple defects in growth and development. Plant Physiol.; 1996; 111, pp. 671-678. [DOI: https://dx.doi.org/10.1104/pp.111.3.671] [PubMed: https://www.ncbi.nlm.nih.gov/pubmed/8754677]

409. Kauschmann, A.; Jessop, A.; Koncz, C.; Szekeres, M.; Willmitzer, L.; Altmann, T. Genetic evidence for an essential role of brassinosteroids in plant development. Plant J.; 1996; 9, pp. 701-713. [DOI: https://dx.doi.org/10.1046/j.1365-313X.1996.9050701.x]

410. Li, J.; Chory, J. A putative leucine-rich repeat receptor kinase involved in brassinosteroid signal transduction. Cell; 1997; 90, pp. 929-938. [DOI: https://dx.doi.org/10.1016/S0092-8674(00)80357-8] [PubMed: https://www.ncbi.nlm.nih.gov/pubmed/9298904]

411. Yamamuro, C.; Ihara, Y.; Wu, X.; Noguchi, T.; Fujioka, S.; Takatsuto, S.; Ashikari, M.; Kitano, H.; Matsuoka, M. Loss of function of a rice brassinosteroid insensitive1 homolog prevents internode elongation and bending of the lamina joint. Plant Cell; 2000; 12, pp. 1591-1606. [DOI: https://dx.doi.org/10.1105/tpc.12.9.1591] [PubMed: https://www.ncbi.nlm.nih.gov/pubmed/11006334]

412. Montoya, T.; Nomura, T.; Farrar, K.; Kaneta, T.; Yokota, T.; Bishop, G.J. Cloning the tomato curl3 gene highlights the putative dual role of the leucine-rich repeat receptor kinase tBRI1/SR160 in plant steroid hormone and peptide hormone signaling. Plant Cell; 2002; 14, pp. 3163-3176. [DOI: https://dx.doi.org/10.1105/tpc.006379]

413. Nomura, T.; Bishop, G.J.; Kaneta, T.; Reid, J.B.; Chory, J.; Yokota, T. The LKA gene is a BRASSINOSTEROID INSENSITIVE 1 homolog of pea. Plant J.; 2003; 36, pp. 291-300. [DOI: https://dx.doi.org/10.1046/j.1365-313X.2003.01863.x] [PubMed: https://www.ncbi.nlm.nih.gov/pubmed/14617087]

414. Chono, M.; Honda, I.; Zeniya, H.; Yoneyama, K.; Saisho, D.; Takeda, K.; Takatsuto, S.; Hoshino, T.; Watanabe, Y. A semidwarf phenotype of barley uzu results from a nucleotide substitution in the gene encoding a putative brassinosteroid receptor. Plant Physiol.; 2003; 133, pp. 1209-1219. [DOI: https://dx.doi.org/10.1104/pp.103.026195] [PubMed: https://www.ncbi.nlm.nih.gov/pubmed/14551335]

415. Holton, N.; Caño-Delgado, A.; Harrison, K.; Montoya, T.; Chory, J.; Bishop, G.J. Tomato BRASSINOSTEROID INSENSITIVE1 is required for systemin-induced root elongation in Solanum pimpinellifolium but is not essential for wound signalling. Plant Cell; 2007; 19, pp. 1709-1717. [DOI: https://dx.doi.org/10.1105/tpc.106.047795]

416. Verhoef, N.; Yokota, T.; Shibata, K.; de Boer, G.J.; Gerats, T.; Vandenbussche, M.; Koes, R.; Souer, E. Brassinosteroid biosynthesis and signalling in Petunia hybrida. J. Exp. Bot.; 2013; 64, pp. 2435-2448. [DOI: https://dx.doi.org/10.1093/jxb/ert102] [PubMed: https://www.ncbi.nlm.nih.gov/pubmed/23599276]

417. Yang, D.H.; Baldwin, I.T.; Wu, J. Silencing brassinosteroid receptor BRI1 impairs herbivory-elicited accumulation of jasmonic acid-isoleucine and diterpene glycosides, but not jasmonic acid and trypsin proteinase inhibitors in Nicotiana attenuata. J. Integr. Plant Biol.; 2013; 55, pp. 514-526. [DOI: https://dx.doi.org/10.1111/jipb.12035] [PubMed: https://www.ncbi.nlm.nih.gov/pubmed/23347255]

418. Feng, Y.; Yin, Y.; Fei, S. Down-regulation of BdBRI1, a putative brassinosteroid receptor gene produces a dwarf phenotype with enhanced drought tolerance in Brachypodium distachyon. Plant Sci.; 2015; 234, pp. 163-173. [DOI: https://dx.doi.org/10.1016/j.plantsci.2015.02.015] [PubMed: https://www.ncbi.nlm.nih.gov/pubmed/25804819]

419. Kir, G.; Ye, H.; Nelissen, H.; Neelakandan, A.K.; Kusnandar, A.S.; Luo, A.; Inzé, D.; Sylvester, A.W.; Yin, Y.; Becraft, P.W. RNA Interference Knockdown of BRASSINOSTEROID INSENSITIVE1 in Maize Reveals Novel Functions for Brassinosteroid Signaling in Controlling Plant Architecture. Plant Physiol.; 2015; 169, pp. 826-839. [DOI: https://dx.doi.org/10.1104/pp.15.00367] [PubMed: https://www.ncbi.nlm.nih.gov/pubmed/26162429]

420. Cheng, X.; Gou, X.; Yin, H.; Mysore, K.S.; Li, J.; Wen, J. Functional characterisation of brassinosteroid receptor MtBRI1 in Medicago truncatula. Sci. Rep.; 2017; 7, 9327. [DOI: https://dx.doi.org/10.1038/s41598-017-09297-9]

421. Caño-Delgado, A.; Yin, Y.; Yu, C.; Vafeados, D.; Mora-García, S.; Cheng, J.C.; Nam, K.H.; Li, J.; Chory, J. BRL1 and BRL3 are novel brassinosteroid receptors that function in vascular differentiation in Arabidopsis. Development; 2004; 131, pp. 5341-5351. [DOI: https://dx.doi.org/10.1242/dev.01403]

422. Wang, X.; Li, X.; Meisenhelder, J.; Hunter, T.; Yoshida, S.; Asami, T.; Chory, J. Autoregulation and homodimerization are involved in the activation of the plant steroid receptor BRI1. Dev. Cell; 2005; 8, pp. 855-865. [DOI: https://dx.doi.org/10.1016/j.devcel.2005.05.001]

423. Wu, G.; Wang, X.; Li, X.; Kamiya, Y.; Otegui, M.S.; Chory, J. Methylation of a phosphatase specifies dephosphorylation and degradation of activated brassinosteroid receptors. Sci. Signal.; 2011; 4, ra29. [DOI: https://dx.doi.org/10.1126/scisignal.2001258] [PubMed: https://www.ncbi.nlm.nih.gov/pubmed/21558554]

424. Wang, R.; Liu, M.; Yuan, M.; Oses-Prieto, J.A.; Cai, X.; Sun, Y.; Burlingame, A.L.; Wang, Z.Y.; Tang, W. The Brassinosteroid-Activated BRI1 Receptor Kinase Is Switched off by Dephosphorylation Mediated by Cytoplasm-Localized PP2A B’ Subunits. Mol. Plant; 2016; 9, pp. 148-157. [DOI: https://dx.doi.org/10.1016/j.molp.2015.10.007] [PubMed: https://www.ncbi.nlm.nih.gov/pubmed/26517938]

425. Wang, X.; Chory, J. Brassinosteroids regulate dissociation of BKI1, a negative regulator of BRI1 signaling, from the plasma membrane. Science; 2006; 313, pp. 1118-1122. [DOI: https://dx.doi.org/10.1126/science.1127593] [PubMed: https://www.ncbi.nlm.nih.gov/pubmed/16857903]

426. Kinoshita, T.; Caño-Delgado, A.; Seto, H.; Hiranuma, S.; Fujioka, S.; Yoshida, S.; Chory, J. Binding of brassinosteroids to the extracellular domain of plant receptor kinase BRI1. Nature; 2005; 433, pp. 167-171. [DOI: https://dx.doi.org/10.1038/nature03227]

427. Wang, X.; Kota, U.; He, K.; Blackburn, K.; Li, J.; Goshe, M.B.; Huber, S.C.; Clouse, S.D. Sequential transphosphorylation of the BRI1/BAK1 receptor kinase complex impacts early events in brassinosteroid signaling. Dev. Cell; 2008; 15, pp. 220-235. [DOI: https://dx.doi.org/10.1016/j.devcel.2008.06.011] [PubMed: https://www.ncbi.nlm.nih.gov/pubmed/18694562]

428. Hothorn, M.; Belkhadir, Y.; Dreux, M.; Dabi, T.; Noel, J.P.; Wilson, I.A.; Chory, J. Structural basis of steroid hormone perception by the receptor kinase BRI1. Nature; 2011; 474, pp. 467-471. [DOI: https://dx.doi.org/10.1038/nature10153]

429. She, J.; Han, Z.; Kim, T.W.; Wang, J.; Cheng, W.; Chang, J.; Shi, S.; Wang, J.; Yang, M.; Wang, Z.Y. et al. Structural insight into brassinosteroid perception by BRI1. Nature; 2011; 474, pp. 472-476. [DOI: https://dx.doi.org/10.1038/nature10178] [PubMed: https://www.ncbi.nlm.nih.gov/pubmed/21666666]

430. Chen, J.; Nolan, T.M.; Ye, H.; Zhang, M.; Tong, H.; Xin, P.; Chu, J.; Chu, C.; Li, Z.; Yin, Y. Arabidopsis WRKY46, WRKY54, and WRKY70 transcription factors are involved in brassinosteroid-regulated plant growth and drought responses. Plant Cell; 2017; 29, pp. 1425-1439. [DOI: https://dx.doi.org/10.1105/tpc.17.00364]

431. Jaillais, Y.; Hothorn, M.; Belkhadir, Y.; Dabi, T.; Nimchuk, Z.L.; Meyerowitz, E.M.; Chory, J. Tyrosine phosphorylation controls brassinosteroid receptor activation by triggering membrane release of its kinase inhibitor. Genes Dev.; 2011; 25, pp. 232-237. [DOI: https://dx.doi.org/10.1101/gad.2001911] [PubMed: https://www.ncbi.nlm.nih.gov/pubmed/21289069]

432. Wang, H.; Yang, C.; Zhang, C.; Wang, N.; Lu, D.; Wang, J.; Zhang, S.; Wang, Z.X.; Ma, H.; Wang, X. Dual role of BKI1 and 14-3-3s in brassinosteroid signaling to link receptor with transcription factors. Dev. Cell; 2011; 21, pp. 825-834. [DOI: https://dx.doi.org/10.1016/j.devcel.2011.08.018]

433. Li, J.; Wen, J.; Lease, K.A.; Doke, J.T.; Tax, F.E.; Walker, J.C. BAK1, an Arabidopsis LRR receptor-like protein kinase, interacts with BRI1 and modulates brassinosteroid signaling. Cell; 2002; 110, pp. 213-222. [DOI: https://dx.doi.org/10.1016/S0092-8674(02)00812-7] [PubMed: https://www.ncbi.nlm.nih.gov/pubmed/12150929]

434. Nam, K.H.; Li, J. BRI1/BAK1, a receptor kinase pair mediating brassinosteroid signaling. Cell; 2002; 110, pp. 203-212. [DOI: https://dx.doi.org/10.1016/S0092-8674(02)00814-0] [PubMed: https://www.ncbi.nlm.nih.gov/pubmed/12150928]

435. Russinova, E.; Borst, J.W.; Kwaaitaal, M.; Caño-Delgado, A.; Yin, Y.; Chory, J.; de Vries, S.C. Heterodimerization and endocytosis of Arabidopsis brassinosteroid receptors BRI1 and AtSERK3 (BAK1). Plant Cell; 2004; 16, pp. 3216-3229. [DOI: https://dx.doi.org/10.1105/tpc.104.025387]

436. Hink, M.A.; Shah, K.; Russinova, E.; de Vries, S.C.; Visser, A.J. Fluorescence fluctuation analysis of Arabidopsis thaliana somatic embryogenesis receptor-like kinase and brassinosteroid insensitive 1 receptor oligomerization. Biophys. J.; 2008; 94, pp. 1052-1062. [DOI: https://dx.doi.org/10.1529/biophysj.107.112003]

437. Wang, X.; Goshe, M.B.; Soderblom, E.J.; Phinney, B.S.; Kuchar, J.A.; Li, J.; Asami, T.; Yoshida, S.; Huber, S.C.; Clouse, S.D. Identification and functional analysis of in vivo phosphorylation sites of the Arabidopsis BRASSINOSTEROID-INSENSITIVE1 receptor kinase. Plant Cell; 2005; 17, pp. 1685-1703. [DOI: https://dx.doi.org/10.1105/tpc.105.031393]

438. Wang, J.; Jiang, J.; Wang, J.; Chen, L.; Fan, S.L.; Wu, J.W.; Wang, X.; Wang, Z.X. Structural insights into the negative regulation of BRI1 signaling by BRI1-interacting protein BKI1. Cell Res.; 2014; 24, pp. 1328-1341. [DOI: https://dx.doi.org/10.1038/cr.2014.132]

439. Karlova, R.; Boeren, S.; Russinova, E.; Aker, J.; Vervoort, J.; de Vries, S.C. The Arabidopsis SOMATIC EMBRYOGENESIS RECEPTOR-LIKE KINASE1 protein complex includes BRASSINOSTEROID-INSENSITIVE1. Plant Cell; 2006; 18, pp. 626-638. [DOI: https://dx.doi.org/10.1105/tpc.105.039412]

440. He, K.; Gou, X.; Yuan, T.; Lin, H.; Asami, T.; Yoshida, S.; Russell, S.D.; Li, J. BAK1 and BKK1 regulate brassinosteroid-dependent growth and brassinosteroid-independent cell-death pathways. Curr. Biol.; 2007; 17, pp. 1109-1115. [DOI: https://dx.doi.org/10.1016/j.cub.2007.05.036] [PubMed: https://www.ncbi.nlm.nih.gov/pubmed/17600708]

441. Muto, H.; Yabe, N.; Asami, T.; Hasunuma, K.; Yamamoto, K.T. Overexpression of constitutive differential growth 1 gene, which encodes a RLCKVII-subfamily protein kinase, causes abnormal differential and elongation growth after organ differentiation in Arabidopsis. Plant Physiol.; 2004; 136, pp. 3124-3133. [DOI: https://dx.doi.org/10.1104/pp.104.046805] [PubMed: https://www.ncbi.nlm.nih.gov/pubmed/15466232]

442. Tang, W.; Kim, T.W.; Oses-Prieto, J.A.; Sun, Y.; Deng, Z.; Zhu, S.; Wang, R.; Burlingame, A.L.; Wang, Z.Y. BSKs mediate signal transduction from the receptor kinase BRI1 in Arabidopsis. Science; 2008; 321, pp. 557-560. [DOI: https://dx.doi.org/10.1126/science.1156973] [PubMed: https://www.ncbi.nlm.nih.gov/pubmed/18653891]

443. Kim, T.W.; Guan, S.; Burlingame, A.L.; Wang, Z.Y. The CDG1 kinase mediates brassinosteroid signal transduction from BRI1 receptor kinase to BSU1 phosphatase and GSK3-like kinase BIN2. Mol. Cell; 2011; 43, pp. 561-571. [DOI: https://dx.doi.org/10.1016/j.molcel.2011.05.037] [PubMed: https://www.ncbi.nlm.nih.gov/pubmed/21855796]

444. Kim, T.W.; Guan, S.; Sun, Y.; Deng, Z.; Tang, W.; Shang, J.X.; Sun, Y.; Burlingame, A.L.; Wang, Z.Y. Brassinosteroid signal transduction from cell-surface receptor kinases to nuclear transcription factors. Nat. Cell Biol.; 2009; 11, pp. 1254-1260. [DOI: https://dx.doi.org/10.1038/ncb1970] [PubMed: https://www.ncbi.nlm.nih.gov/pubmed/19734888]

445. Mora-García, S.; Vert, G.; Yin, Y.; Caño-Delgado, A.; Cheong, H.; Chory, J. Nuclear protein phosphatases with Kelch-repeat domains modulate the response to brassinosteroids in Arabidopsis. Genes Dev.; 2004; 18, pp. 448-460. [DOI: https://dx.doi.org/10.1101/gad.1174204] [PubMed: https://www.ncbi.nlm.nih.gov/pubmed/14977918]

446. Li, J.; Nam, K.H. Regulation of brassinosteroid signaling by a GSK3/SHAGGY-like kinase. Science; 2002; 295, pp. 1299-1301. [DOI: https://dx.doi.org/10.1126/science.1065769] [PubMed: https://www.ncbi.nlm.nih.gov/pubmed/11847343]

447. Maselli, G.A.; Slamovits, C.H.; Bianchi, J.I.; Vilarrasa-Blasi, J.; Caño-Delgado, A.I.; Mora-García, S. Revisiting the evolutionary history and roles of protein phosphatases with Kelch-like domains in plants. Plant Physiol.; 2014; 164, pp. 1527-1541. [DOI: https://dx.doi.org/10.1104/pp.113.233627]

448. Yan, Z.; Zhao, J.; Peng, P.; Chihara, R.K.; Li, J. BIN2 functions redundantly with other Arabidopsis GSK3-like kinases to regulate brassinosteroid signaling. Plant Physiol.; 2009; 150, pp. 710-721. [DOI: https://dx.doi.org/10.1104/pp.109.138099] [PubMed: https://www.ncbi.nlm.nih.gov/pubmed/19395409]

449. Peng, P.; Yan, Z.; Zhu, Y.; Li, J. Regulation of the Arabidopsis GSK3-like kinase BRASSINOSTEROID-INSENSITIVE 2 through proteasome-mediated protein degradation. Mol. Plant; 2008; 1, pp. 338-346. [DOI: https://dx.doi.org/10.1093/mp/ssn001]

450. Zhu, J.Y.; Li, Y.; Cao, D.M.; Yang, H.; Oh, E.; Bi, Y.; Zhu, S.; Wang, Z.Y. The F-box Protein KIB1 Mediates Brassinosteroid-Induced Inactivation and Degradation of GSK3-like Kinases in Arabidopsis. Mol. Cell; 2017; 66, pp. 648-657.e4. [DOI: https://dx.doi.org/10.1016/j.molcel.2017.05.012] [PubMed: https://www.ncbi.nlm.nih.gov/pubmed/28575660]

451. Yin, Y.; Wang, Z.Y.; Mora-García, S.; Li, J.; Yoshida, S.; Asami, T.; Chory, J. BES1 accumulates in the nucleus in response to brassinosteroids to regulate gene expression and promote stem elongation. Cell; 2002; 109, pp. 181-191. [DOI: https://dx.doi.org/10.1016/S0092-8674(02)00721-3]

452. Wang, Z.Y.; Nakano, T.; Gendron, J.; He, J.; Chen, M.; Vafeados, D.; Yang, Y.; Fujioka, S.; Yoshida, S.; Asami, T. et al. Nuclear-localized BZR1 mediates brassinosteroid-induced growth and feedback suppression of brassinosteroid biosynthesis. Dev. Cell; 2002; 2, pp. 505-513. [DOI: https://dx.doi.org/10.1016/S1534-5807(02)00153-3] [PubMed: https://www.ncbi.nlm.nih.gov/pubmed/11970900]

453. He, J.X.; Gendron, J.M.; Yang, Y.; Li, J.; Wang, Z.Y. The GSK3-like kinase BIN2 phosphorylates and destabilizes BZR1, a positive regulator of the brassinosteroid signaling pathway in Arabidopsis. Proc. Natl. Acad. Sci. USA; 2002; 99, pp. 10185-10190. [DOI: https://dx.doi.org/10.1073/pnas.152342599] [PubMed: https://www.ncbi.nlm.nih.gov/pubmed/12114546]

454. Vert, G.; Chory, J. Downstream nuclear events in brassinosteroid signalling. Nature; 2006; 441, pp. 96-100. [DOI: https://dx.doi.org/10.1038/nature04681] [PubMed: https://www.ncbi.nlm.nih.gov/pubmed/16672972]

455. Gampala, S.S.; Kim, T.W.; He, J.X.; Tang, W.; Deng, Z.; Bai, M.Y.; Guan, S.; Lalonde, S.; Sun, Y.; Gendron, J.M. et al. An essential role for 14-3-3 proteins in brassinosteroid signal transduction in Arabidopsis. Dev. Cell; 2007; 13, pp. 177-189. [DOI: https://dx.doi.org/10.1016/j.devcel.2007.06.009] [PubMed: https://www.ncbi.nlm.nih.gov/pubmed/17681130]

456. Bai, M.Y.; Zhang, L.Y.; Gampala, S.S.; Zhu, S.W.; Song, W.Y.; Chong, K.; Wang, Z.Y. Functions of OsBZR1 and 14-3-3 proteins in brassinosteroid signaling in rice. Proc. Natl. Acad. Sci. USA; 2007; 104, pp. 13839-13844. [DOI: https://dx.doi.org/10.1073/pnas.0706386104] [PubMed: https://www.ncbi.nlm.nih.gov/pubmed/17699623]

457. Tang, W.; Yuan, M.; Wang, R.; Yang, Y.; Wang, C.; Oses-Prieto, J.A.; Kim, T.W.; Zhou, H.W.; Deng, Z.; Gampala, S.S. et al. PP2A activates brassinosteroid-responsive gene expression and plant growth by dephosphorylating BZR1. Nat. Cell Biol.; 2011; 13, pp. 124-131. [DOI: https://dx.doi.org/10.1038/ncb2151] [PubMed: https://www.ncbi.nlm.nih.gov/pubmed/21258370]

458. Yin, Y.; Vafeados, D.; Tao, Y.; Yoshida, S.; Asami, T.; Chory, J. A new class of transcription factors mediates brassinosteroid-regulated gene expression in Arabidopsis. Cell; 2005; 120, pp. 249-259. [DOI: https://dx.doi.org/10.1016/j.cell.2004.11.044]

459. He, J.X.; Gendron, J.M.; Sun, Y.; Gampala, S.S.; Gendron, N.; Sun, C.Q.; Wang, Z.Y. BZR1 is a transcriptional repressor with dual roles in brassinosteroid homeostasis and growth responses. Science; 2005; 307, pp. 1634-1638. [DOI: https://dx.doi.org/10.1126/science.1107580]

460. Sun, Y.; Fan, X.Y.; Cao, D.M.; Tang, W.; He, K.; Zhu, J.Y.; He, J.X.; Bai, M.Y.; Zhu, S.; Oh, E. et al. Integration of brassinosteroid signal transduction with the transcription network for plant growth regulation in Arabidopsis. Dev. Cell; 2010; 19, pp. 765-777. [DOI: https://dx.doi.org/10.1016/j.devcel.2010.10.010] [PubMed: https://www.ncbi.nlm.nih.gov/pubmed/21074725]

461. Yu, X.; Li, L.; Zola, J.; Aluru, M.; Ye, H.; Foudree, A.; Guo, H.; Anderson, S.; Aluru, S.; Liu, P. et al. A brassinosteroid transcriptional network revealed by genome-wide identification of BES1 target genes in Arabidopsis thaliana. Plant J.; 2011; 65, pp. 634-646. [DOI: https://dx.doi.org/10.1111/j.1365-313X.2010.04449.x] [PubMed: https://www.ncbi.nlm.nih.gov/pubmed/21214652]

462. Nemhauser, J.L.; Mockler, T.C.; Chory, J. Interdependency of brassinosteroid and auxin signaling in Arabidopsis. PLoS Biol.; 2004; 2, E258. [DOI: https://dx.doi.org/10.1371/journal.pbio.0020258]

463. Carretero-Paulet, L.; Galstyan, A.; Roig-Villanova, I.; Martínez-García, J.F.; Bilbao-Castro, J.R.; Robertson, D.L. Genome-wide classification and evolutionary analysis of the bHLH family of transcription factors in Arabidopsis, poplar, rice, moss, and algae. Plant Physiol.; 2010; 153, pp. 1398-1412. [DOI: https://dx.doi.org/10.1104/pp.110.153593]

464. Friedrichsen, D.M.; Nemhauser, J.; Muramitsu, T.; Maloof, J.N.; Alonso, J.; Ecker, J.R.; Furuya, M.; Chory, J. Three redundant brassinosteroid early response genes encode putative bHLH transcription factors required for normal growth. Genetics; 2002; 162, pp. 1445-1456. [DOI: https://dx.doi.org/10.1093/genetics/162.3.1445]

465. Zhou, J.; Liu, D.; Wang, P.; Ma, X.; Lin, W.; Chen, S.; Mishev, K.; Lu, D.; Kumar, R.; Vanhoutte, I. et al. Regulation of Arabidopsis brassinosteroid receptor BRI1 endocytosis and degradation by plant U-box PUB12/PUB13-mediated ubiquitination. Proc. Natl. Acad. Sci. USA; 2018; 115, pp. E1906-E1915. [DOI: https://dx.doi.org/10.1073/pnas.1712251115] [PubMed: https://www.ncbi.nlm.nih.gov/pubmed/29432171]

466. Naranjo-Arcos, M.; Srivastava, M.; Deligne, F.; Bhagat, P.K.; Mansi, M.; Sadanandom, A.; Vert, G. SUMO/deSUMOylation of the BRI1 brassinosteroid receptor modulates plant growth responses to temperature. Proc. Natl. Acad. Sci. USA; 2023; 120, e2217255120. [DOI: https://dx.doi.org/10.1073/pnas.2217255120] [PubMed: https://www.ncbi.nlm.nih.gov/pubmed/36652487]

467. Anwar, A.; Liu, Y.; Dong, R.; Bai, L.; Yu, X.; Li, Y. The physiological and molecular mechanism of brassinosteroid in response to stress: A review. Biol. Res.; 2018; 51, 46. [DOI: https://dx.doi.org/10.1186/s40659-018-0195-2]

468. Singh, J.; Nakamura, S.; Ota, Y. Effect of epi-brassinolide on gram plants grown under water-stress in juvenile stage. Indian J. Agric. Sci.; 1993; 63, pp. 395-397. [DOI: https://dx.doi.org/10.1002/jsfa.2740610404]

469. Ye, H.; Liu, S.; Tang, B.; Chen, J.; Xie, Z.; Nolan, T.M.; Jiang, H.; Guo, H.; Lin, H.Y.; Li, L. et al. RD26 mediates crosstalk between drought and brassinosteroid signalling pathways. Nat. Commun.; 2017; 8, 14573. [DOI: https://dx.doi.org/10.1038/ncomms14573]

470. Fàbregas, N.; Lozano-Elena, F.; Blasco-Escámez, D.; Tohge, T.; Martínez-Andújar, C.; Albacete, A.; Osorio, S.; Bustamante, M.; Riechmann, J.L.; Nomura, T. et al. Overexpression of the vascular brassinosteroid receptor BRL3 confers drought resistance without penalizing plant growth. Nat. Commun.; 2018; 9, 4680. [DOI: https://dx.doi.org/10.1038/s41467-018-06861-3]

471. Nie, S.; Huang, S.; Wang, S.; Mao, Y.; Liu, J.; Ma, R.; Wang, X. Enhanced brassinosteroid signaling intensity via SlBRI1 overexpression negatively regulates drought resistance in a manner opposite of that via exogenous BR application in tomato. Plant Physiol. Biochem.; 2019; 138, pp. 36-47. [DOI: https://dx.doi.org/10.1016/j.plaphy.2019.02.014] [PubMed: https://www.ncbi.nlm.nih.gov/pubmed/30844693]

472. Xiang, Y.; Li, G.; Li, Q.; Niu, Y.; Pan, Y.; Cheng, Y.; Bian, X.; Zhao, C.; Wang, Y.; Zhang, A. Autophagy receptor ZmNBR1 promotes the autophagic degradation of ZmBRI1a and enhances drought tolerance in maize. J. Integr. Plant Biol.; 2024; 66, pp. 1068-1086. [DOI: https://dx.doi.org/10.1111/jipb.13662] [PubMed: https://www.ncbi.nlm.nih.gov/pubmed/38607264]

473. Koh, S.; Lee, S.C.; Kim, M.K.; Koh, J.H.; Lee, S.; An, G.; Choe, S.; Kim, S.R. T-DNA tagged knockout mutation of rice OsGSK1, an orthologue of Arabidopsis BIN2, with enhanced tolerance to various abiotic stresses. Plant Mol. Biol.; 2007; 65, pp. 453-466. [DOI: https://dx.doi.org/10.1007/s11103-007-9213-4]

474. Guo, X.; Zhang, J.; Sun, S.; Huang, L.; Niu, Y.; Zhao, P.; Zhang, Y.; Shi, X.; Ji, W.; Xu, S. TaGSK3 regulates wheat development and stress adaptation through BR-dependent and BR-independent pathways. Plant Cell Environ.; 2024; 47, pp. 2443-2458. [DOI: https://dx.doi.org/10.1111/pce.14890]

475. Cai, Z.; Liu, J.; Wang, H.; Yang, C.; Chen, Y.; Li, Y.; Pan, S.; Dong, R.; Tang, G.; de Dios Barajas-Lopez, J. et al. GSK3-like kinases positively modulate abscisic acid signaling through phosphorylating subgroup III SnRK2s in Arabidopsis. Proc. Natl. Acad. Sci. USA; 2014; 111, pp. 9651-9656. [DOI: https://dx.doi.org/10.1073/pnas.1316717111]

476. Wang, H.; Tang, J.; Liu, J.; Hu, J.; Liu, J.; Chen, Y.; Cai, Z.; Wang, X. Abscisic acid signaling inhibits brassinosteroid signaling through dampening the dephosphorylation of BIN2 by ABI1 and ABI2. Mol. Plant; 2018; 11, pp. 315-325. [DOI: https://dx.doi.org/10.1016/j.molp.2017.12.013]

477. Xie, Z.; Nolan, T.; Jiang, H.; Tang, B.; Zhang, M.; Li, Z.; Yin, Y. The AP2/ERF transcription factor TINY modulates brassinosteroid-regulated plant growth and drought responses in Arabidopsis. Plant Cell; 2019; 31, pp. 1788-1806. [DOI: https://dx.doi.org/10.1105/tpc.18.00918] [PubMed: https://www.ncbi.nlm.nih.gov/pubmed/31126980]

478. Van Nguyen, T.; Park, C.R.; Lee, K.H.; Lee, S.; Kim, C.S. BES1/BZR1 homolog 3 cooperates with E3 ligase AtRZF1 to regulate osmotic stress and brassinosteroid responses in Arabidopsis. J. Exp. Bot.; 2021; 72, pp. 636-653. [DOI: https://dx.doi.org/10.1093/jxb/eraa458] [PubMed: https://www.ncbi.nlm.nih.gov/pubmed/33529338]

479. Park, C.R.; Nguyen, V.T.; Min, J.H.; Sang, H.; Lim, G.H.; Kim, C.S. Isolation and functional characterization of soybean BES1/BZR1 homolog 3-like 1 (GmBEH3L1) associated with dehydration sensitivity and brassinosteroid signaling in Arabidopsis thaliana. Plants; 2022; 11, 2565. [DOI: https://dx.doi.org/10.3390/plants11192565] [PubMed: https://www.ncbi.nlm.nih.gov/pubmed/36235431]

480. Cui, X.Y.; Gao, Y.; Guo, J.; Yu, T.F.; Zheng, W.J.; Liu, Y.W.; Chen, J.; Xu, Z.S.; Ma, Y.Z. BES/BZR transcription factor TaBZR2 positively regulates drought responses by activation of TaGST1. Plant Physiol.; 2019; 180, pp. 605-620. [DOI: https://dx.doi.org/10.1104/pp.19.00100]

481. Surina, S.; Yamagami, A.; Miyaji, T.; Chagan, Z.; Chung, K.M.; Mitsuda, N.; Nishida, K.; Tachibana, R.; Zhu, Z.; Miyakawa, T. et al. BIL9 promotes both plant growth via BR signaling and drought stress resistance by binding with the transcription factor HDG11. Plant Cell Physiol.; 2024; 65, pcae009. [DOI: https://dx.doi.org/10.1093/pcp/pcae009]

482. Nie, W.F.; Wang, M.M.; Xia, X.J.; Zhou, Y.H.; Shi, K.; Chen, Z.; Yu, J.Q. Silencing of tomato RBOH1 and MPK2 abolishes brassinosteroid-induced H₂O₂ generation and stress tolerance. Plant Cell Environ.; 2013; 36, pp. 789-803. [DOI: https://dx.doi.org/10.1111/pce.12014]

483. Zhou, J.; Wang, J.; Li, X.; Xia, X.J.; Zhou, Y.H.; Shi, K.; Chen, Z.; Yu, J.Q. H₂O₂ mediates the crosstalk of brassinosteroid and abscisic acid in tomato responses to heat and oxidative stresses. J. Exp. Bot.; 2014; 65, pp. 4371-4383. [DOI: https://dx.doi.org/10.1093/jxb/eru217] [PubMed: https://www.ncbi.nlm.nih.gov/pubmed/24899077]

484. Xia, X.J.; Gao, C.J.; Song, L.X.; Zhou, Y.H.; Shi, K.; Yu, J.Q. Role of H₂O₂ dynamics in brassinosteroid-induced stomatal closure and opening in Solanum lycopersicum. Plant Cell Environ.; 2014; 37, pp. 2036-2050. [DOI: https://dx.doi.org/10.1111/pce.12275]

485. Zhu, T.; Deng, X.; Zhou, X.; Zhu, L.; Zou, L.; Li, P.; Zhang, D.; Lin, H. Ethylene and hydrogen peroxide are involved in brassinosteroid-induced salt tolerance in tomato. Sci. Rep.; 2016; 6, 35392. [DOI: https://dx.doi.org/10.1038/srep35392]

486. Bender, K.W.; Wang, X.; Cheng, G.B.; Kim, H.S.; Zielinski, R.E.; Huber, S.C. Glutaredoxin AtGRXC2 catalyses inhibitory glutathionylation of Arabidopsis BRI1-associated receptor-like kinase 1 (BAK1) in vitro. Biochem. J.; 2015; 467, pp. 399-413. [DOI: https://dx.doi.org/10.1042/BJ20141403] [PubMed: https://www.ncbi.nlm.nih.gov/pubmed/25678081]

487. Tian, Y.; Fan, M.; Qin, Z.; Lv, H.; Wang, M.; Zhang, Z.; Zhou, W.; Zhao, N.; Li, X.; Han, C. et al. Hydrogen peroxide positively regulates brassinosteroid signaling through oxidation of the BRASSINAZOLE-RESISTANT1 transcription factor. Nat. Commun.; 2018; 9, 1063. [DOI: https://dx.doi.org/10.1038/s41467-018-03463-x]

488. Samanta, S.; Seth, C.S.; Roychoudhury, A. The molecular paradigm of reactive oxygen species (ROS) and reactive nitrogen species (RNS) with different phytohormone signaling pathways during drought stress in plants. Plant Physiol. Biochem.; 2024; 206, 108259. [DOI: https://dx.doi.org/10.1016/j.plaphy.2023.108259] [PubMed: https://www.ncbi.nlm.nih.gov/pubmed/38154293]

489. Umehara, M.; Hanada, A.; Yoshida, S.; Akiyama, K.; Arite, T.; Takeda-Kamiya, N.; Magome, H.; Kamiya, Y.; Shirasu, K.; Yoneyama, K. et al. Inhibition of shoot branching by new terpenoid plant hormones. Nature; 2008; 455, pp. 195-200. [DOI: https://dx.doi.org/10.1038/nature07272] [PubMed: https://www.ncbi.nlm.nih.gov/pubmed/18690207]

490. Gomez-Roldan, V.; Fermas, S.; Brewer, P.B.; Puech-Pagès, V.; Dun, E.A.; Pillot, J.P.; Letisse, F.; Matusova, R.; Danoun, S.; Portais, J.C. et al. Strigolactone inhibition of shoot branching. Nature; 2008; 455, pp. 189-194. [DOI: https://dx.doi.org/10.1038/nature07271] [PubMed: https://www.ncbi.nlm.nih.gov/pubmed/18690209]

491. Cook, C.E.; Whichard, L.P.; Turner, B.; Wall, M.E.; Egley, G.H. Germination of Witchweed (Striga lutea Lour.): Isolation and Properties of a Potent Stimulant. Science; 1966; 154, pp. 1189-1190. [DOI: https://dx.doi.org/10.1126/science.154.3753.1189] [PubMed: https://www.ncbi.nlm.nih.gov/pubmed/17780042]

492. Al-Babili, S.; Bouwmeester, H.J. Strigolactones, a novel carotenoid-derived plant hormone. Annu. Rev. Plant Biol.; 2015; 66, pp. 161-186. [DOI: https://dx.doi.org/10.1146/annurev-arplant-043014-114759]

493. de Saint Germain, A.; Clavé, G.; Badet-Denisot, M.A.; Pillot, J.P.; Cornu, D.; Le Caer, J.P.; Burger, M.; Pelissier, F.; Retailleau, P.; Turnbull, C. et al. An histidine covalent receptor and butenolide complex mediates strigolactone perception. Nat. Chem. Biol.; 2016; 12, pp. 787-794. [DOI: https://dx.doi.org/10.1038/nchembio.2147]

494. Snowden, K.C.; Simkin, A.J.; Janssen, B.J.; Templeton, K.R.; Loucas, H.M.; Simons, J.L.; Karunairetnam, S.; Gleave, A.P.; Clark, D.G.; Klee, H.J. The Decreased apical dominance1/Petunia hybrida CAROTENOID CLEAVAGE DIOXYGENASE8 gene affects branch production and plays a role in leaf senescence, root growth, and flower development. Plant Cell; 2005; 17, pp. 746-759. [DOI: https://dx.doi.org/10.1105/tpc.104.027714]

495. Scaffidi, A.; Waters, M.T.; Ghisalberti, E.L.; Dixon, K.W.; Flematti, G.R.; Smith, S.M. Carlactone-independent seedling morphogenesis in Arabidopsis. Plant J.; 2013; 76, pp. 1-9. [DOI: https://dx.doi.org/10.1111/tpj.12265] [PubMed: https://www.ncbi.nlm.nih.gov/pubmed/23773129]

496. Stirnberg, P.; van De Sande, K.; Leyser, H.M. MAX1 and MAX2 control shoot lateral branching in Arabidopsis. Development; 2002; 129, pp. 1131-1141. [DOI: https://dx.doi.org/10.1242/dev.129.5.1131]

497. Yamada, Y.; Furusawa, S.; Nagasaka, S.; Shimomura, K.; Yamaguchi, S.; Umehara, M. Strigolactone signaling regulates rice leaf senescence in response to a phosphate deficiency. Planta; 2014; 240, pp. 399-408. [DOI: https://dx.doi.org/10.1007/s00425-014-2096-0]

498. Sang, D.; Chen, D.; Liu, G.; Liang, Y.; Huang, L.; Meng, X.; Chu, J.; Sun, X.; Dong, G.; Yuan, Y. et al. Strigolactones regulate rice tiller angle by attenuating shoot gravitropism through inhibiting auxin biosynthesis. Proc. Natl. Acad. Sci. USA; 2014; 111, pp. 11199-11204. [DOI: https://dx.doi.org/10.1073/pnas.1411859111] [PubMed: https://www.ncbi.nlm.nih.gov/pubmed/25028496]

499. Agusti, J.; Herold, S.; Schwarz, M.; Sanchez, P.; Ljung, K.; Dun, E.A.; Brewer, P.B.; Beveridge, C.A.; Sieberer, T.; Sehr, E.M. et al. Strigolactone signaling is required for auxin-dependent stimulation of secondary growth in plants. Proc. Natl. Acad. Sci. USA; 2011; 108, pp. 20242-20247. [DOI: https://dx.doi.org/10.1073/pnas.1111902108] [PubMed: https://www.ncbi.nlm.nih.gov/pubmed/22123958]

500. Toh, S.; Kamiya, Y.; Kawakami, N.; Nambara, E.; McCourt, P.; Tsuchiya, Y. Thermoinhibition Uncovers a Role for Strigolactones in Arabidopsis Seed Germination. Plant Cell Physiol.; 2012; 53, pp. 107-117. [DOI: https://dx.doi.org/10.1093/pcp/pcr176] [PubMed: https://www.ncbi.nlm.nih.gov/pubmed/22173099]

501. Tsuchiya, Y.; Vidaurre, D.; Toh, S.; Hanada, A.; Nambara, E.; Kamiya, Y.; Yamaguchi, S.; McCourt, P. A Small-Molecule Screen Identifies New Functions for the Plant Hormone Strigolactone. Nat. Chem. Biol.; 2010; 6, pp. 741-749. [DOI: https://dx.doi.org/10.1038/nchembio.435] [PubMed: https://www.ncbi.nlm.nih.gov/pubmed/20818397]

502. Arite, T.; Kameoka, H.; Kyozuka, J. Strigolactone Positively Controls Crown Root Elongation in Rice. J. Plant Growth Regul.; 2012; 31, pp. 165-172. [DOI: https://dx.doi.org/10.1007/s00344-011-9228-6]

503. Kagiyama, M.; Hirano, Y.; Mori, T.; Kim, S.Y.; Kyozuka, J.; Seto, Y.; Yamaguchi, S.; Hakoshima, T. Structures of D14 and D14L in the Strigolactone and Karrikin Signaling Pathways. Genes Cells; 2013; 18, pp. 147-160. [DOI: https://dx.doi.org/10.1111/gtc.12025] [PubMed: https://www.ncbi.nlm.nih.gov/pubmed/23301669]

504. Kapulnik, Y.; Delaux, P.M.; Resnick, N.; Mayzlish-Gati, E.; Wininger, S.; Bhattacharya, C.; Séjalon-Delmas, N.; Combier, J.P.; Bécard, G.; Belausov, E. et al. Strigolactones Affect Lateral Root Formation and Root-Hair Elongation in Arabidopsis. Planta; 2011; 233, pp. 209-216. [DOI: https://dx.doi.org/10.1007/s00425-010-1310-y]

505. Kohlen, W.; Charnikhova, T.; Lammers, M.; Pollina, T.; Tóth, P.; Haider, I.; Pozo, M.J.; de Maagd, R.A.; Ruyter-Spira, C.; Bouwmeester, H.J. et al. The Tomato CAROTENOID CLEAVAGE DIOXYGENASE8 (SlCCD8) Regulates Rhizosphere Signaling, Plant Architecture, and Affects Reproductive Development Through Strigolactone Biosynthesis. New Phytol.; 2012; 196, pp. 535-547. [DOI: https://dx.doi.org/10.1111/j.1469-8137.2012.04265.x]

506. Rasmussen, A.; Depuydt, S.; Goormachtig, S.; Geelen, D. Strigolactones Fine-Tune the Root System. Planta; 2013; 238, pp. 615-626. [DOI: https://dx.doi.org/10.1007/s00425-013-1911-3] [PubMed: https://www.ncbi.nlm.nih.gov/pubmed/23801297]

507. Urquhart, S.; Foo, E.; Reid, J.B. The Role of Strigolactones in Photomorphogenesis of Pea is Limited to Adventitious Rooting. Physiol. Plant; 2015; 153, pp. 392-402. [DOI: https://dx.doi.org/10.1111/ppl.12246]

508. Brewer, P.B.; Koltai, H.; Beveridge, C.A. Diverse roles of strigolactones in plant development. Mol. Plant; 2013; 6, pp. 18-28. [DOI: https://dx.doi.org/10.1093/mp/sss130] [PubMed: https://www.ncbi.nlm.nih.gov/pubmed/23155045]

509. Ruyter-Spira, C.; Al-Babili, S.; van der Krol, S.; Bouwmeester, H. The biology of strigolactones. Trends Plant Sci.; 2013; 18, pp. 72-83. [DOI: https://dx.doi.org/10.1016/j.tplants.2012.10.003] [PubMed: https://www.ncbi.nlm.nih.gov/pubmed/23182342]

510. Sun, H.; Tao, J.; Gu, P.; Xu, G.; Zhang, Y. The role of strigolactones in root development. Plant Signal. Behav.; 2016; 11, e1110662. [DOI: https://dx.doi.org/10.1080/15592324.2015.1110662] [PubMed: https://www.ncbi.nlm.nih.gov/pubmed/26515106]

511. Waldie, T.; McCulloch, H.; Leyser, O. Strigolactones and the control of plant development: Lessons from shoot branching. Plant J.; 2014; 79, pp. 607-622. [DOI: https://dx.doi.org/10.1111/tpj.12488] [PubMed: https://www.ncbi.nlm.nih.gov/pubmed/24612082]

512. Zwanenburg, B.; Pospíšil, T.; Ćavar Zeljković, S. Strigolactones: New plant hormones in action. Planta; 2016; 243, pp. 1311-1326. [DOI: https://dx.doi.org/10.1007/s00425-015-2455-5] [PubMed: https://www.ncbi.nlm.nih.gov/pubmed/26838034]

513. Alder, A.; Jamil, M.; Marzorati, M.; Bruno, M.; Vermathen, M.; Bigler, P.; Ghisla, S.; Bouwmeester, H.; Beyer, P.; Al-Babili, S. The path from β-carotene to carlactone, a strigolactone-like plant hormone. Science; 2012; 335, pp. 1348-1351. [DOI: https://dx.doi.org/10.1126/science.1218094]

514. Kretzschmar, T.; Kohlen, W.; Sasse, J.; Borghi, L.; Schlegel, M.; Bachelier, J.B.; Reinhardt, D.; Bours, R.; Bouwmeester, H.J.; Martinoia, E. A petunia ABC protein controls strigolactone-dependent symbiotic signalling and branching. Nature; 2012; 483, pp. 341-344. [DOI: https://dx.doi.org/10.1038/nature10873] [PubMed: https://www.ncbi.nlm.nih.gov/pubmed/22398443]

515. Kang, J.; Hwang, J.U.; Lee, M.; Kim, Y.Y.; Assmann, S.M.; Martinoia, E.; Lee, Y. PDR-type ABC transporter mediates cellular uptake of the phytohormone abscisic acid. Proc. Natl. Acad. Sci. USA; 2010; 107, pp. 2355-2360. [DOI: https://dx.doi.org/10.1073/pnas.0909222107] [PubMed: https://www.ncbi.nlm.nih.gov/pubmed/20133880]

516. Kuromori, T.; Miyaji, T.; Yabuuchi, H.; Shimizu, H.; Sugimoto, E.; Kamiya, A.; Moriyama, Y.; Shinozaki, K. ABC transporter AtABCG25 is involved in abscisic acid transport and responses. Proc. Natl. Acad. Sci. USA; 2010; 107, pp. 2361-2366. [DOI: https://dx.doi.org/10.1073/pnas.0912516107]

517. Yao, R.; Ming, Z.; Yan, L.; Li, S.; Wang, F.; Ma, S.; Yu, C.; Yang, M.; Chen, L.; Chen, L. et al. DWARF14 is a non-canonical hormone receptor for strigolactone. Nature; 2016; 536, pp. 469-473. [DOI: https://dx.doi.org/10.1038/nature19073] [PubMed: https://www.ncbi.nlm.nih.gov/pubmed/27479325]

518. Jiang, L.; Liu, X.; Xiong, G.; Liu, H.; Chen, F.; Wang, L.; Meng, X.; Liu, G.; Yu, H.; Yuan, Y. et al. DWARF 53 acts as a repressor of strigolactone signalling in rice. Nature; 2013; 504, pp. 401-405. [DOI: https://dx.doi.org/10.1038/nature12870]

519. Smith, S.M.; Li, J. Signalling and responses to strigolactones and karrikins. Curr. Opin. Plant Biol.; 2014; 21, pp. 23-29. [DOI: https://dx.doi.org/10.1016/j.pbi.2014.06.003] [PubMed: https://www.ncbi.nlm.nih.gov/pubmed/24996032]

520. Xiong, G.; Wang, Y.; Li, J. Action of strigolactones in plants. Enzymes; 2014; 35, pp. 57-84. [DOI: https://dx.doi.org/10.1016/B978-0-12-801922-1.00003-8] [PubMed: https://www.ncbi.nlm.nih.gov/pubmed/25740715]

521. Stanga, J.P.; Morffy, N.; Nelson, D.C. Functional redundancy in the control of seedling growth by the karrikin signaling pathway. Planta; 2016; 243, pp. 1397-1406. [DOI: https://dx.doi.org/10.1007/s00425-015-2458-2]

522. Duan, J.; Yu, H.; Yuan, K.; Liao, Z.; Meng, X.; Jing, Y.; Liu, G.; Chu, J.; Li, J. Strigolactone promotes cytokinin degradation through transcriptional activation of CYTOKININ OXIDASE/DEHYDROGENASE 9 in rice. Proc. Natl. Acad. Sci. USA; 2019; 116, pp. 14319-14324. [DOI: https://dx.doi.org/10.1073/pnas.1810980116] [PubMed: https://www.ncbi.nlm.nih.gov/pubmed/31235564]

523. Huang, X.; Hilscher, J.; Stoger, E.; Christou, P.; Zhu, C. Modification of cereal plant architecture by genome editing to improve yields. Plant Cell Rep.; 2021; 40, pp. 953-978. [DOI: https://dx.doi.org/10.1007/s00299-021-02668-7]

524. Okazaki, K.; Watanabe, S.; Koike, I.; Kawada, K.; Ito, S.; Nakamura, H.; Asami, T.; Shimomura, K.; Umehara, M. Strigolactone signaling inhibition increases adventitious shoot formation on internodal segments of ipecac. Planta; 2021; 253, 123. [DOI: https://dx.doi.org/10.1007/s00425-021-03640-1] [PubMed: https://www.ncbi.nlm.nih.gov/pubmed/34014387]

525. Zhang, J.; Mazur, E.; Balla, J.; Gallei, M.; Kalousek, P.; Medveďová, Z.; Li, Y.; Wang, Y.; Prát, T.; Vasileva, M. et al. Strigolactones inhibit auxin feedback on PIN-dependent auxin transport canalization. Nature Commun.; 2020; 11, 3508. [DOI: https://dx.doi.org/10.1038/s41467-020-17252-y] [PubMed: https://www.ncbi.nlm.nih.gov/pubmed/32665554]

526. Asghar, S.; Xiong, Y.; Che, M.; Fan, X.; Li, H.; Wang, Y.; Xu, X.; Li, W.; Han, Z. Transcriptome analysis reveals the effects of strigolactone on shoot regeneration of apple. Plant Cell Rep.; 2022; 41, pp. 1613-1626. [DOI: https://dx.doi.org/10.1007/s00299-022-02882-x] [PubMed: https://www.ncbi.nlm.nih.gov/pubmed/35680714]

527. Ha, C.V.; Leyva-González, M.A.; Osakabe, Y.; Tran, U.T.; Nishiyama, R.; Watanabe, Y.; Tanaka, M.; Seki, M.; Yamaguchi, S.; Dong, N.V. et al. Positive regulatory role of strigolactone in plant responses to drought and salt stress. Proc. Natl. Acad. Sci. USA; 2014; 111, pp. 851-856. [DOI: https://dx.doi.org/10.1073/pnas.1322135111]

528. Nisa, Z.U.; Wang, Y.; Ali, N.; Chen, C.; Zhang, X.; Jin, X.; Yu, L.; Jing, L.; Chen, C.; Elansary, H.O. Strigolactone signaling gene from soybean GmMAX2a enhances the drought and salt-alkaline resistance in Arabidopsis via regulating transcriptional profiles of stress-related genes. Funct. Integr. Genom.; 2023; 23, 216. [DOI: https://dx.doi.org/10.1007/s10142-023-01151-8] [PubMed: https://www.ncbi.nlm.nih.gov/pubmed/37391642]

529. Mostofa, M.G.; Rahman, M.M.; Nguyen, K.H.; Li, W.; Watanabe, Y.; Tran, C.D.; Zhang, M.; Itouga, M.; Fujita, M.; Tran, L.P. Strigolactones regulate arsenate uptake, vacuolar-sequestration and antioxidant defense responses to resist arsenic toxicity in rice roots. J. Hazard. Mater.; 2021; 415, 125589. [DOI: https://dx.doi.org/10.1016/j.jhazmat.2021.125589]

530. Vurro, M.; Prandi, C.; Baroccio, F. Strigolactones: How far is their commercial use for agricultural purposes?. Pest Manag. Sci.; 2016; 72, pp. 2026-2034. [DOI: https://dx.doi.org/10.1002/ps.4254] [PubMed: https://www.ncbi.nlm.nih.gov/pubmed/26869010]

531. Carreon-Gonzalez, M.; Alvarez-Idaboy, J.R. The Synergy between Glutathione and Phenols-Phenolic Antioxidants Repair Glutathione: Closing the Virtuous Circle-A Theoretical Insight. Antioxidants; 2023; 12, 1125. [DOI: https://dx.doi.org/10.3390/antiox12051125]

532. Tai, Z.; Yin, X.; Fang, Z.; Shi, G.; Lou, L.; Cai, Q. Exogenous GR24 Alleviates Cadmium Toxicity by Reducing Cadmium Uptake in Switchgrass (Panicum virgatum) Seedlings. Int. J. Environ. Res. Public Health; 2017; 14, 852. [DOI: https://dx.doi.org/10.3390/ijerph14080852] [PubMed: https://www.ncbi.nlm.nih.gov/pubmed/28758909]

533. Singh, A.; Roychoudhury, A. Abscisic acid in plants under abiotic stress: Crosstalk with major phytohormones. Plant Cell Rep.; 2023; 42, pp. 961-974. [DOI: https://dx.doi.org/10.1007/s00299-023-03013-w]

534. Jogawat, A. Crosstalk among phytohormone signaling pathways during abiotic stress. Molecular Plant Abiotic Stress: Biology and Biotechnology; Roychoudhury, A.; Tripathi, D. Wiley: Hoboken, NJ, USA, 2019; pp. 209-220.

535. Müller, M.; Munné-Bosch, S. Hormonal impact on photosynthesis and photoprotection in plants. Plant Physiol.; 2021; 185, pp. 1500-1522. [DOI: https://dx.doi.org/10.1093/plphys/kiaa119]

536. Desikan, R.; Last, K.; Harrett-Williams, R.; Tagliavia, C.; Harter, K.; Hooley, R.; Hancock, J.T.; Neill, S.J. Ethylene-induced stomatal closure in Arabidopsis occurs via AtrbohF-mediated hydrogen peroxide synthesis. Plant J.; 2006; 47, pp. 907-916. [DOI: https://dx.doi.org/10.1111/j.1365-313X.2006.02842.x]

537. Cheng, M.C.; Liao, P.M.; Kuo, W.W.; Lin, T.P. The Arabidopsis ETHYLENE RESPONSE FACTOR1 regulates abiotic stress-responsive gene expression by binding to different cis-acting elements in response to different stress signals. Plant Physiol.; 2013; 162, pp. 1566-1582. [DOI: https://dx.doi.org/10.1104/pp.113.221911]

538. Tanaka, Y.; Sano, T.; Tamaoki, M.; Nakajima, N.; Kondo, N.; Hasezawa, S. Ethylene inhibits abscisic acid induced stomatal closure in Arabidopsis. Plant Physiol.; 2005; 138, pp. 2337-2343. [DOI: https://dx.doi.org/10.1104/pp.105.063503] [PubMed: https://www.ncbi.nlm.nih.gov/pubmed/16024687]

539. Watkins, J.M.; Hechler, P.J.; Muday, G.K. Ethylene induced favonol accumulation in guard cells suppresses reactive oxygen species and moderates stomatal aperture. Plant Physiol.; 2014; 164, pp. 1707-1717. [DOI: https://dx.doi.org/10.1104/pp.113.233528] [PubMed: https://www.ncbi.nlm.nih.gov/pubmed/24596331]

540. Watkins, J.M.; Chapman, J.M.; Muday, G.K. Abscisic acid-induced reactive oxygen species are modulated by favonols to control stomata aperture. Plant Physiol.; 2017; 175, pp. 1807-1825. [DOI: https://dx.doi.org/10.1104/pp.17.01010]

541. Rasheed, F.; Sehar, Z.; Fatma, M.; Iqbal, N.; Masood, A.; Anjum, N.A.; Khan, N.A. Involvement of ethylene in reversal of salt stress by salicylic acid in the presence of sulfur in mustard (Brassica juncea L.). J. Plant Growth Regul.; 2021; 41, pp. 3449-3466. [DOI: https://dx.doi.org/10.1007/s00344-021-10526-9]

542. Fatma, M.; Iqbal, N.; Gautam, H.; Sehar, Z.; Sofo, A.; D’Ippolito, I.; Khan, N.A. Ethylene and sulfur coordinately modulate the antioxidant system and ABA accumulation in mustard plants under salt stress. Plants; 2021; 10, 180. [DOI: https://dx.doi.org/10.3390/plants10010180] [PubMed: https://www.ncbi.nlm.nih.gov/pubmed/33478097]

543. Poór, P.; Nawaz, K.; Gupta, R.; Ashfaque, F.; Khan, M.I. Ethylene involvement in the regulation of heat stress tolerance in plants. Plant Rep.; 2021; 41, pp. 675-698. [DOI: https://dx.doi.org/10.1007/s00299-021-02675-8]

544. McMichael, B.L.; Jordan, W.R.; Powell, R.D. An effect of water stress on ethylene production by intact cotton petioles. Plant Physiol.; 1972; 49, pp. 658-660. [DOI: https://dx.doi.org/10.1104/pp.49.4.658] [PubMed: https://www.ncbi.nlm.nih.gov/pubmed/16658022]

545. Dong, Z.; Yu, Y.; Li, S.; Wang, J.; Tang, S.; Huang, R. Abscisic acid antagonizes ethylene production through the ABI4-mediated transcriptional repression of ACS4 and ACS8 in Arabidopsis. Mol. Plant; 2016; 9, pp. 126-135. [DOI: https://dx.doi.org/10.1016/j.molp.2015.09.007] [PubMed: https://www.ncbi.nlm.nih.gov/pubmed/26410794]

546. Voisin, A.S.; Reidy, B.; Parent, B.; Rolland, G.; Redondo, E.; Gerentes, D.; Tardieu, F.; Muller, B. Are ABA, ethylene or their interaction involved in the response of leaf growth to soil water deficit? An analysis using naturally occurring variation or genetic transformation of ABA production in maize. Plant Cell Environ.; 2006; 29, pp. 1829-1840. [DOI: https://dx.doi.org/10.1111/j.1365-3040.2006.01560.x]

547. Sharp, R.E.; LeNoble, M.E.; Else, M.A.; Thorne, E.T.; Gherardi, F. Endogenous ABA maintains shoot growth in tomato independently of effects on plant water balance: Evidence for an interaction with ethylene. J. Exp. Bot.; 2000; 51, pp. 1575-1584. [DOI: https://dx.doi.org/10.1093/jexbot/51.350.1575] [PubMed: https://www.ncbi.nlm.nih.gov/pubmed/11006308]

548. Nazareno, A.L.; Hernandez, B.S. A mathematical model of the interaction of abscisic acid, ethylene and methyl jasmonate on stomatal closure in plants. PLoS ONE; 2017; 12, e0171065. [DOI: https://dx.doi.org/10.1371/journal.pone.0171065] [PubMed: https://www.ncbi.nlm.nih.gov/pubmed/28182683]

549. Habben, J.E.; Bao, X.; Bate, N.J.; DeBruin, J.L.; Dolan, D.; Hasegawa, D.; Helentjaris, T.G.; Lafitte, R.H.; Lovan, N.; Mo, H. et al. Transgenic alteration of ethylene biosynthesis increases grain yield in maize under field drought-stress conditions. Plant Biotechnol. J.; 2014; 12, pp. 685-693. [DOI: https://dx.doi.org/10.1111/pbi.12172] [PubMed: https://www.ncbi.nlm.nih.gov/pubmed/24618117]

550. Tholen, D.; Pons, T.L.; Voesenek, L.A.C.J.; Poorter, H. The role of ethylene perception in the control of photosynthesis. Plant Signal. Behav.; 2008; 3, pp. 108-109. [DOI: https://dx.doi.org/10.4161/psb.3.2.4968] [PubMed: https://www.ncbi.nlm.nih.gov/pubmed/19704724]

551. Da Silva, J.M.; Arrabaça, M.C. Photosynthesis in the water-stressed C4 grass Setaria sphacelata is mainly limited by stomata with both rapidly and slowly imposed water deficits. Physiol. Plant.; 2004; 121, pp. 409-420. [DOI: https://dx.doi.org/10.1111/j.1399-3054.2004.00328.x]

552. Xu, Z.; Zhou, G.; Shimizu, H. Are plant growth and photosynthesis limited by pre-drought following rewatering in grass?. J. Exp. Bot.; 2009; 60, pp. 3737-3749. [DOI: https://dx.doi.org/10.1093/jxb/erp216]

553. Jardim-Messeder, D.; Caverzan, A.; Rauber, R.; Cunha, J.R.; Carvalho, F.E.L.; Gaeta, M.L.; Da Fonseca, G.C.; Costa, J.M.; Frei, M.; Silveira, J.A.G. et al. Thylakoidal APX modulates hydrogen peroxide content and stomatal closure in rice (Oryza sativa L.). Environ. Exp. Bot.; 2018; 150, pp. 46-56. [DOI: https://dx.doi.org/10.1016/j.envexpbot.2018.02.012]

554. Xu, W.; Jia, L.; Shi, W.; Liang, J.; Zhou, F.; Li, Q.; Zhang, J. Abscisic acid accumulation modulates auxin transport in the root tip to enhance proton secretion for maintaining root growth under moderate water stress. New Phytol.; 2013; 197, pp. 139-150. [DOI: https://dx.doi.org/10.1111/nph.12004]

555. Xie, Q.; Essemine, J.; Pang, X.; Chen, H.; Cai, W. Exogenous application of abscisic acid to shoots promotes primary root cell division and elongation. Plant Sci.; 2020; 292, 110385. [DOI: https://dx.doi.org/10.1016/j.plantsci.2019.110385]

556. Zhang, Y.; Wang, X.; Luo, Y.; Zhang, L.; Yao, Y.; Han, L.; Chen, Z.; Wang, L.; Li, Y. OsABA8ox2, an ABA catabolic gene, suppresses root elongation of rice seedlings and contributes to drought response. Crop J.; 2020; 8, pp. 480-491. [DOI: https://dx.doi.org/10.1016/j.cj.2019.08.006]

557. Zhang, Y.; Du, H.; Gui, Y.; Xu, F.; Liu, J.; Zhang, J.; Xu, W. Moderate water stress in rice induces rhizosheath formation associated with abscisic acid and auxin responses. J. Exp. Bot.; 2021; 71, pp. 2740-2751. [DOI: https://dx.doi.org/10.1093/jxb/eraa021] [PubMed: https://www.ncbi.nlm.nih.gov/pubmed/32053723]

558. Miao, R.; Yuan, W.; Wang, Y.; Garcia-Maquilon, I.; Dang, X.; Li, Y.; Zhang, J.; Zhu, Y.; Rodriguez, P.L.; Xu, W. Low ABA concentration promotes root growth and hydrotropism through relief of ABA INSENSITIVE 1-mediated inhibition of plasma membrane H+-ATPase 2. Sci. Adv.; 2021; 7, eabd4113. [DOI: https://dx.doi.org/10.1126/sciadv.abd4113]

559. Aslam, M.; Sugano, S.S.; Takahashi, H. Hydrotropism in plants: Mechanisms and biological significance. Plant Cell Environ.; 2022; 45, pp. 315-327. [DOI: https://dx.doi.org/10.1111/pce.14123]

560. Qin, H.; Wang, J.; Zhou, J.; Qiao, J.; Li, Y.; Quan, R.; Huang, R. Abscisic acid promotes auxin biosynthesis to inhibit primary root elongation in rice. Plant Physiol.; 2023; 191, pp. 1953-1967. [DOI: https://dx.doi.org/10.1093/plphys/kiac586] [PubMed: https://www.ncbi.nlm.nih.gov/pubmed/36535001]

561. Blum, A. Plant Breeding for Water-Limited Environments; Springer: New York, NY, USA, 2011; pp. 217-234.

562. Umezawa, T.; Nakashima, K.; Miyakawa, T.; Kuromori, T.; Tanokura, M.; Shinozaki, K.; Yamaguchi-Shinozaki, K. Molecular basis of the core regulatory network in ABA responses: Sensing, signaling and transport. Plant Cell Physiol.; 2010; 51, pp. 1821-1839. [DOI: https://dx.doi.org/10.1093/pcp/pcq156] [PubMed: https://www.ncbi.nlm.nih.gov/pubmed/20980270]

563. Wang, H.; Wang, X. GSK3-like Kinases Are a Class of Positive Components in the Core ABA Signaling Pathway. Mol. Plant; 2018; 11, pp. 761-763. [DOI: https://dx.doi.org/10.1016/j.molp.2018.03.019]

564. Tang, X.; Wang, C.; Liu, Y.; He, G.; Ma, N.; Chai, G.; Li, S.; Xu, H.; Zhou, G. Brassinosteroid Signaling Converges With Auxin-Mediated C3H17 to Regulate Xylem Formation in Populus. Front. Plant Sci.; 2020; 11, 586014. [DOI: https://dx.doi.org/10.3389/fpls.2020.586014] [PubMed: https://www.ncbi.nlm.nih.gov/pubmed/33193536]

565. Zheng, B.; Bai, Q.; Wu, L.; Liu, H.; Liu, Y.; Xu, W.; Li, G.; Ren, H.; She, X.; Wu, G. EMS1 and BRI1 control separate biological processes via extracellular domain diversity and intracellular domain conservation. Nat. Commun.; 2019; 10, 4165. [DOI: https://dx.doi.org/10.1038/s41467-019-12112-w] [PubMed: https://www.ncbi.nlm.nih.gov/pubmed/31519884]

566. Chen, W.; Lv, M.; Wang, Y.; Wang, P.A.; Cui, Y.; Li, M.; Wang, R.; Gou, X.; Li, J. BES1 is activated by EMS1-TPD1-SERK1/2-mediated signaling to control tapetum development in Arabidopsis thaliana. Nat. Commun.; 2019; 10, 4164. [DOI: https://dx.doi.org/10.1038/s41467-019-12118-4]

567. Cai, H.; Huang, Y.; Liu, L.; Zhang, M.; Chai, M.; Xi, X.; Aslam, M.; Wang, L.; Ma, S.; Su, H. et al. Signaling by the EPFL-ERECTA family coordinates female germline specification through the BZR1 family in Arabidopsis. Plant Cell; 2023; 35, pp. 1455-1473. [DOI: https://dx.doi.org/10.1093/plcell/koad032] [PubMed: https://www.ncbi.nlm.nih.gov/pubmed/36748257]

568. Cao, X.; Wei, Y.; Shen, B.; Liu, L.; Mao, J. Interaction of the Transcription Factors BES1/BZR1 in Plant Growth and Stress Response. Int. J. Mol. Sci.; 2024; 25, 6836. [DOI: https://dx.doi.org/10.3390/ijms25136836] [PubMed: https://www.ncbi.nlm.nih.gov/pubmed/38999944]

569. Gallego-Bartolomé, J.; Minguet, E.G.; Grau-Enguix, F.; Abbas, M.; Locascio, A.; Thomas, S.G.; Alabadí, D.; Blázquez, M.A. Molecular mechanism for the interaction between gibberellin and brassinosteroid signaling pathways in Arabidopsis. Proc. Natl. Acad. Sci. USA; 2012; 109, pp. 13446-13451. [DOI: https://dx.doi.org/10.1073/pnas.1119992109]

570. Li, Q.F.; Wang, C.; Jiang, L.; Li, S.; Sun, S.S.; He, J.X. An interaction between BZR1 and DELLAs mediates direct signaling crosstalk between brassinosteroids and gibberellins in Arabidopsis. Sci. Signal.; 2012; 5, ra72. [DOI: https://dx.doi.org/10.1126/scisignal.2002908]

571. Oh, E.; Zhu, J.Y.; Wang, Z.Y. Interaction between BZR1 and PIF4 integrates brassinosteroid and environmental responses. Nat. Cell Biol.; 2012; 14, pp. 802-809. [DOI: https://dx.doi.org/10.1038/ncb2545]

572. De Ollas, C.; Hernando, B.; Arbona, V.; Gómez-Cadenas, A. Jasmonic acid transient accumulation is needed for abscisic acid increase in citrus roots under drought stress conditions. Physiol. Plant.; 2013; 147, pp. 296-306. [DOI: https://dx.doi.org/10.1111/j.1399-3054.2012.01659.x] [PubMed: https://www.ncbi.nlm.nih.gov/pubmed/22671923]

573. Jang, G.; Chang, S.H.; Um, T.Y.; Lee, S.; Kim, J.K.; Choi, Y. Do Antagonistic interaction between jasmonic acid and cytokinin in xylem development. Sci. Rep.; 2017; 7, 10212. [DOI: https://dx.doi.org/10.1038/s41598-017-10634-1]

574. Chen, Q.; Sun, J.; Zhai, Q.; Zhou, W.; Qi, L.; Xu, L.; Wang, B.; Chen, R.; Jiang, H.; Qi, J. et al. The basic helix-loop-helix transcription factor myc2 directly represses plethora expression during jasmonate-mediated modulation of the root stem cell niche in Arabidopsis. Plant Cell; 2011; 23, pp. 3335-3352. [DOI: https://dx.doi.org/10.1105/tpc.111.089870]

575. Ren, C.; Han, C.; Peng, W.; Huang, Y.; Peng, Z.; Xiong, X.; Zhu, Q.; Gao, B.; Xie, D. A leaky mutation in DWARF4 reveals an antagonistic role of brassinosteroid in the inhibition of root growth by jasmonate in Arabidopsis. Plant Physiol.; 2009; 151, pp. 1412-1420. [DOI: https://dx.doi.org/10.1104/pp.109.140202]

576. Nahar, K.; Kyndt, T.; Hause, B.; Höfte, M.; Gheysen, G. Brassinosteroids suppress rice defense against root-knot nematodes through antagonism with the jasmonate pathway. Mol. Plant. Microbe Interact.; 2013; 26, pp. 106-115. [DOI: https://dx.doi.org/10.1094/MPMI-05-12-0108-FI] [PubMed: https://www.ncbi.nlm.nih.gov/pubmed/23194343]

577. Liu, X.; Hou, X. Antagonistic Regulation of ABA and GA in Metabolism and Signaling Pathways. Front. Plant Sci.; 2018; 9, 251. [DOI: https://dx.doi.org/10.3389/fpls.2018.00251] [PubMed: https://www.ncbi.nlm.nih.gov/pubmed/29535756]

578. Oh, E.; Yamaguchi, S.; Hu, J.H.; Yusuke, J.; Jung, B.; Paik, I.; Lee, H.S.; Sun, T.P.; Kamiya, Y.; Choi, G. PIL5, a phytochrome-interacting bHLH protein, regulates gibberellin responsiveness by binding directly to the GAI and RGA promoters in Arabidopsis seeds. Plant Cell; 2007; 19, pp. 1192-1208. [DOI: https://dx.doi.org/10.1105/tpc.107.050153]

579. Seo, M.; Hanada, A.; Kuwahara, A.; Endo, A.; Okamoto, M.; Yamauchi, Y.; Nambara, E. Regulation of hormone metabolism in Arabidopsis seeds: Phytochrome regulation of abscisic acid metabolism and abscisic acid regulation of gibberellin metabolism. Plant J.; 2006; 48, pp. 354-366. [DOI: https://dx.doi.org/10.1111/j.1365-313X.2006.02881.x] [PubMed: https://www.ncbi.nlm.nih.gov/pubmed/17010113]

580. Du, H.; Chang, Y.; Huang, F.; Xiong, L. GID1 modulates stomatal response and submergence tolerance involving abscisic acid and gibberellic acid signaling in rice. J. Integr. Plant Biol.; 2015; 57, pp. 954-968. [DOI: https://dx.doi.org/10.1111/jipb.12313] [PubMed: https://www.ncbi.nlm.nih.gov/pubmed/25418692]

581. Finkelstein, R.R.; Lynch, T.J. Overexpression of ABI5 Binding Proteins Suppresses Inhibition of Germination Due to Overaccumulation of DELLA Proteins. Int. J. Mol. Sci.; 2022; 23, 5537. [DOI: https://dx.doi.org/10.3390/ijms23105537] [PubMed: https://www.ncbi.nlm.nih.gov/pubmed/35628355]

582. Hou, X.; Lee, L.Y.; Xia, K.; Yan, Y.; Yu, H. DELLAs modulate jasmonate signaling via competitive binding to JAZs. Dev. Cell; 2010; 19, pp. 884-894. [DOI: https://dx.doi.org/10.1016/j.devcel.2010.10.024]

583. Cao, D.; Cheng, H.; Wu, W.; Soo, H.M.; Peng, J. Gibberellin mobilizes distinct DELLA dependent transcriptomes to regulate seed germination and floral development in Arabidopsis. Plant Physiol.; 2006; 142, pp. 509-525. [DOI: https://dx.doi.org/10.1104/pp.106.082289]

584. Hou, X.; Hu, W.W.; Shen, L.; Lee, L.Y.; Tao, Z.; Han, J.H.; Yu, H. Global identification of DELLA target genes during Arabidopsis flower development. Plant Physiol.; 2008; 147, pp. 1126-1142. [DOI: https://dx.doi.org/10.1104/pp.108.121301]

585. Navarro, L.; Bari, R.; Achard, P.; Lison, P.; Nemri, A.; Harberd, N.P.; Jones, J.D. DELLAs control plant immune responses by modulating the balance of jasmonic acid and salicylic acid signaling. Curr. Biol.; 2008; 18, pp. 650-655. [DOI: https://dx.doi.org/10.1016/j.cub.2008.03.060] [PubMed: https://www.ncbi.nlm.nih.gov/pubmed/18450451]

586. Yuan, H.; Zhao, L.; Guo, W.; Yu, Y.; Tao, L.; Zhang, L.; Song, X.; Huang, W.; Cheng, L.; Chen, J. et al. Exogenous application of phytohormones promotes growth and regulates expression of wood formation-related genes in Populus simonii × P. nigra. Int. J. Mol. Sci.; 2019; 20, 792. [DOI: https://dx.doi.org/10.3390/ijms20030792]

587. Willige, B.C.; Isono, E.; Richter, R.; Zourelidou, M.; Schwechheimer, C. Gibberellin regulates PIN-FORMED abundance and is required for auxin transport-dependent growth and development in Arabidopsis thaliana. Plant Cell; 2011; 23, pp. 2184-2195. [DOI: https://dx.doi.org/10.1105/tpc.111.086355] [PubMed: https://www.ncbi.nlm.nih.gov/pubmed/21642547]

588. Castro-Camba, R.; Sánchez, C.; Vidal, N.; Vielba, J.M. Interactions of Gibberellins with Phytohormones and Their Role in Stress Responses. Horticulturae; 2022; 8, 241. [DOI: https://dx.doi.org/10.3390/horticulturae8030241]

589. Gallego-Bartolomé, J.; Arana, M.V.; Vandenbussche, F.; Žádníková, P.; Minguet, E.G.; Guardiola, V.; Van Der Straeten, D.; Benkova, E.; Alabadí, D.; Blázquez, M.A. Hierarchy of hormone action controlling apical hook development in Arabidopsis. Plant J.; 2011; 67, pp. 622-634. [DOI: https://dx.doi.org/10.1111/j.1365-313X.2011.04621.x]

590. Li, W.; Herrera-Estrella, L.; Tran, L.P. The Yin-Yang of Cytokinin Homeostasis and Drought Acclimation/Adaptation. Trends Plant Sci.; 2016; 21, pp. 548-550. [DOI: https://dx.doi.org/10.1016/j.tplants.2016.05.006]

591. Corot, A.; Roman, H.; Douillet, O.; Autret, H.; Perez-Garcia, M.D.; Citerne, S.; Bertheloot, J.; Sakr, S.; Leduc, N.; Demotes-Mainard, S. Cytokinins and Abscisic Acid Act Antagonistically in the Regulation of the Bud Outgrowth Pattern by Light Intensity. Front. Plant Sci.; 2017; 8, 1724. [DOI: https://dx.doi.org/10.3389/fpls.2017.01724]

592. Davies, W.J.; Kudoyarova, G.; Hartung, W. Long-Distance ABA Signaling and Its Relation to Other Signaling Pathways in the Detection of Soil Drying and the Mediation of the Plant’s Response to Drought. J. Plant Growth Regul.; 2005; 24, pp. 285-295. [DOI: https://dx.doi.org/10.1007/s00344-005-0103-1]

593. Huang, X.; Hou, L.; Meng, J.; You, H.; Li, Z.; Gong, Z.; Yang, S.; Shi, Y. The Antagonistic Action of Abscisic Acid and Cytokinin Signaling Mediates Drought Stress Response in Arabidopsis. Mol. Plant; 2018; 11, pp. 970-982. [DOI: https://dx.doi.org/10.1016/j.molp.2018.05.001] [PubMed: https://www.ncbi.nlm.nih.gov/pubmed/29753021]

594. Jeon, J.; Kim, N.Y.; Kim, S.; Kang, N.Y.; Novák, O.; Ku, S.J.; Cho, C.; Lee, D.J.; Lee, E.J.; Strnad, M. et al. A Subset of Cytokinin Two-Component Signaling System Plays a Role in Cold Temperature Stress Response in Arabidopsis. J. Biol. Chem.; 2010; 285, pp. 23371-23386. [DOI: https://dx.doi.org/10.1074/jbc.M109.096644] [PubMed: https://www.ncbi.nlm.nih.gov/pubmed/20463025]

595. Wang, Y.; Li, L.; Ye, T.; Zhao, S.; Liu, Z.; Feng, Y.Q.; Wu, Y. Cytokinin Antagonizes ABA Suppression to Seed Germination of Arabidopsis by Downregulating ABI5 Expression. Plant J.; 2011; 68, pp. 249-261. [DOI: https://dx.doi.org/10.1111/j.1365-313X.2011.04683.x]

596. Huang, X.; Zhang, X.; Gong, Z.; Yang, S.; Shi, Y. ABI4 Represses the Expression of Type-A ARRs to Inhibit Seed Germination in Arabidopsis. Plant J.; 2017; 89, pp. 354-365. [DOI: https://dx.doi.org/10.1111/tpj.13389] [PubMed: https://www.ncbi.nlm.nih.gov/pubmed/27711992]

597. Tran, L.S.; Shinozaki, K.; Yamaguchi-Shinozaki, K. Role of Cytokinin Responsive Two-Component System in ABA and Osmotic Stress Signaling. Plant Signal. Behav.; 2010; 5, pp. 148-150. [DOI: https://dx.doi.org/10.4161/psb.5.2.10411]

598. Tran, L.S.; Urao, T.; Qin, F.; Maruyama, K.; Kakimoto, T.; Shinozaki, K.; Yamaguchi-Shinozaki, K. Functional Analysis of AHK1/ATHK1 and Cytokinin Receptor Histidine Kinases in Response to Abscisic Acid, Drought, and Salt Stress in Arabidopsis. Proc. Natl. Acad. Sci. USA; 2007; 104, pp. 20623-20628. [DOI: https://dx.doi.org/10.1073/pnas.0706547105] [PubMed: https://www.ncbi.nlm.nih.gov/pubmed/18077346]