Introduction

Strigolactones (SLs) comprise a novel class of phytohormones first discovered as germination inducers of various parasitic plant species (Cook et al. 1966; Kohlen et al. 2011). Their name originates from their role in stimulating Striga (parasitic witchweeds) germination and from their characteristic lactone ring structure. The first isolated Striga seed germination inducers were strigyl acetate and strigol from Gossypium hirsutum L. (Cook et al. 1966). Retrospectively, SLs were first indicated as phytohormones through their presence as unknown graft-transmissible signals that suppressed Pisum sativum shoot branching (Beveridge et al. 1994). Signal-deficient mutants showed a hyper branching phenotype that was independent of known phytohormones, like cytokinins and auxins (Koltai 2014).

Two research groups then independently identified SLs as new phytohormones regulating the shoot branching phenotypes (Gomez-Roldan et al. 2008; Umehara et al. 2008). Plant shoot branching is inhibited by endogenous SL production or exogenous SL application in these hyper branching mutants (Umehara et al. 2008) (Fig. 1). Root and shoot extracts of various species, including Arabidopsis, contain various types, combinations, and levels of SL molecules (Goldwasser et al. 2008; Koltai and Beveridge 2013; Kapulnik and Koltai. 2014; Saeed et al. 2017; Bürger and Chory 2020). To regulate shoot branching, root-derived SLs are mainly transported to shoots through the xylem (Kohlen et al. 2011; Borghi et al. 2016). Since the discovery of SLs as phytohormones, extensive research has revealed novel insights about their diversity, biosynthesis, and signaling. Because of their important roles in plant growth and development, SLs can potentially be used for crop improvement. For example, mutating the SL biosynthetic gene HTD1/D17 increases rice yields, which contributed to the “Green Revolution” since the 1960s (Wang et al. 2020a).

Fig. 1
figure 1

Diverse roles of SLs in overall plant growth, development, and resilience

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SLs are characterized by their butenolide moieties – lactones with a 4-C heterocyclic ring structure (Omoarelojie et al. 2019). These hormones are at the forefront of plant science research because of their diverse biological roles, ranging from growth and development to interactions with other organisms (Agusti et al. 2011; Cook et al. 1966; Toh et al. 2012; Domagalska and Leyser 2011). The synthetic SL analog GR24 is an important tool in investigating the functions of SLs in plant physiology (Arite et al. 2009). It has been most useful in species without known SL biosynthetic/signaling mutants and its application reverses SL biosynthetic but not signaling mutant phenotypes (Gomez-Roldan et al. 2008; Umehara et al. 2008).

Although initially considered to be detrimental to plants since they enhanced parasitic plant germination (Cook et al. 1966), SLs were later considered beneficial since they also mediate arbuscular mycorrhizal (AM) fungal colonization (Akiyama et al. 2005; Besserer et al. 2006). Moreover, they initiate AM fungal hyphal branching even before host root infection (Akiyama et al. 2005). SLs also interact with rhizobia and affect nodule formation in leguminous plants, reflecting their diverse roles in biotic interactions (Foo et al. 2014). Apart from their functions in regulating plant symbiotic relationships, SLs may mediate defenses against pathogens (Torres-Vera et al. 2014).

In addition, SLs can effectively alleviate various abiotic stresses (Fig. 1), such as salt and drought stresses (Ma et al. 2017; Van Ha et al. 2014; Lu et al. 2019). In Arabidopsis thaliana, SLs can regulate adaptive responses, such as stress-induced changes in stomatal density and closure (Van Ha et al. 2014). In their study, SL-deficient plants were hypersensitive to such stresses (Van Ha et al. 2014). Exogenous SL application rescued drought-sensitive mutant phenotypes, while it augmented the drought tolerance of wild-type (WT) plants (Van Ha et al. 2014).

Other hormones interact with SLs to regulate various physiological processes, enabling plants to respond to changing environmental factors, such as nutrient availability, shading, and temperature (Cheng et al. 2013). For example, auxins work together with SLs to control shoot branching patterns (Hayward et al. 2009; Bennett et al. 2016; Ligerot et al. 2017). SLs and abscisic acid (ABA) work together during abiotic stresses (Ren et al. 2018). Moreover, ethylene and SLs act antagonistically to control hypocotyl growth (Yu et al. 2013).

Strigolactone Biosynthesis: From Humble Pigment Beginnings

SLs and SL-like compounds have a conserved lactone structure consisting of three rings (ABC rings) connected through an enol ether bridge with a fourth methyl butenolide or furanone moiety (D ring) (Al-Babili and Bouwmeester 2015; Yoneyama et al. 2018). The region connecting the core (ABC) with the D ring acts as the bioactiphore (Zwanenburg et al. 2009). Endogenous SLs are classified into two main types (strigol and orobanchol type) based on whether the C ring is α- or β-oriented (Cui. 2014). Strigol and orobanchol are canonical SLs as both have A, B, C, and D rings (Butler. 1995); around 23 types of canonical SLs have been characterized in root exudates (Xie et al. 2010). Certain SL-like compounds are considered non-canonical, because they lack the A, B, and/or C ring; however, they still possess the D ring bonded to the rest of the molecule (Alder et al. 2012; Boyer et al. 2014; Waters et al. 2017). Non-canonical SLs include certain synthetic and natural compounds like methyl carlactonoate (MeCLA), avenaol, and Yoshimulactone Green (Abe et al. 2014; Kim et al. 2014; Tsuchiya et al. 2015). The structural diversity in canonical SLs stems from various AB ring system modifications, including epoxidation, hydroxylation, ketolation, and oxidation (Bhattacharya et al. 2009). This wide structural diversity involves many SL biosynthetic genes (Saeed et al. 2017), homologs of which have been found in algae and bryophytes (Delaux et al. 2012).

Several studies have elucidated the molecular mechanism of SL biosynthesis. The involvement of the carotenoid pathway was reported using fluridone, an inhibitor of carotenoid biosynthesis (Matusova et al. 2005). SL biosynthesis has also been investigated using certain carotenoid catabolic mutants (Matusova et al. 2005), and different branching mutants such as P. sativum ramosus (rms) mutants (Johnson et al. 2006; Beveridge et al. 1994), Arabidopsis max (more axillary growth) mutants (Sorefan et al. 2003) and Petunia decreased apical dominance (dad1, dad2, dad3) mutants (Snowden et al. 2005). Gene cloning, reciprocal grafting experiments and mutant analysis implied that SLs are synthesized from carotenoids and are transported acropetally (Ongaro et al. 2008).

SL biosynthesis initially occurs in the chloroplasts (Alder et al. 2012; Saeed et al. 2017) involving DWARF27 (D27/β-carotene isomerase), which requires iron as a cofactor (Lin et al. 2009). D27 catalyzes β-carotene isomerization by acting on its 9th chemical bond, changing its configuration from trans-β-carotene into 9-cis-β-carotene (C-40) (Alder et al. 2012). These carotenoids have a 40-carbon skeleton with an extended conjugated double bond system (Moise et al. 2014). Downstream of D27, carotenoid cleavage dioxygenases (CCDs) convert carotenoids into apocarotenoids (Auldridge et al. 2006; Waters et al., 2012a; Hou et al. 2016), which are then modified by other CCD enzymes (Alder et al. 2008). Oxidation of various carotenoid precursors, resulting in specific double bond breakage, yields various compounds like ABA, SLs, and retinal (a conjugated chromophore) (Felemban et al. 2019). The Arabidopsis genome encodes about nine different CCDs (CCD1-9), five of which are 9-cis-epoxycarotenoid cleavage dioxygenase (NCEDs) involved in ABA biosynthesis (Tan et al. 2003). In addition, various enzymes encoded by MAX genes (MAX1, MAX3 and MAX4) regulate SL biosynthesis in Arabidopsis (Ruyter-Spira et al. 2013). ABA itself may also regulate SL biosynthesis, because ABA-deficient maize (vp14) and tomato (notabilis) mutants showed lower seed germination (Matusova et al. 2005).

In molecular detail, CCD-catalyzed SL biosynthesis produces intermediates that are further oxidized by cytochrome P450s (Matusova et al. 2005). Two known CCDs (CCD7 and CCD8) act progressively in the pathway; CCD7 is encoded by MAX3 and its orthologs RMS5 and D17/HTD1 (Booker et al. 2004), whereas CCD8 is encoded by MAX4 and its orthologs RMS1, D10, and DAD1 (Arite et al. 2007). 9-cis-β-carotene is converted by CCD7 into 9-cis-β-apo-10-carotenal (C-27) and β ionone (C-13) (Waters et al. 2012a). 9-cis-β-apo-10-carotenal is then converted by CCD8 into Carlactone (CL), a possible mobile intermediate containing two rings (A and D) along with the enol ether bridge and an SL-like carbon skeleton (Alder et al. 2012; Seto et al. 2014). CL is produced by intra-molecular rearrangement of 9-cis-β-apo-10-carotenal, which suggests that each β-carotene molecule produces a single SL molecule (Alder et al. 2012; Seto et al. 2014). CL has similar properties as SLs, such as stimulating seed germination of Striga hermonthica, and is a putative intermediate during the biosynthesis of other SLs (Alder et al. 2012). Seto and colleagues (2014) used 13C-labeled CL to detect its conversion into SLs in vivo. Conversion of exogenous CL into SL has been reported in rice, suggesting that CL is the precursor of endogenous SLs (Seto et al. 2014). Remarkably, Baz et al. (2018) reported that a new product 3-OH-carlactone is formed in vitro from 9-cis-3-OH-β-apo-10′-carotenal by the action of D27, CCD7 and CCD8. They also showed 3-OH-carlactone formation in planta by expressing rice and Arabidopsis CL biosynthetic genes in Nicotiana benthamiana leaves (Baz et al. 2018).

CL is subsequently transported into the cytoplasm for further processing (Al-Babili and Bouwmeester 2015). CL (with a complete D ring) acts as the common precursor of all SLs; however, it needs further modifications since it lacks the B and C rings (Alder et al. 2012). CL is then converted into carlactonoic acid (CLA) by the cytochrome P450 monooxygenase enzyme MAX1 in Arabidopsis (Abe et al. 2014; Zhang et al. 2014). Booker et al. (2005) demonstrated the role of MAX1 (CYP711A1) in CLA synthesis, by reciprocal grafting experiments in A. thaliana. In these experiments, the excessive branching phenotype of max4 (ccd8) mutant scions were eventually reversed by grafting with wild-type MAX1 root stocks (Booker et al. 2005). The conversion of CL into CLA in vitro using recombinant MAX1 protein inside yeast microsomes further clarified the function of MAX1 (Abe et al. 2014). MAX1 catalyzes back-to-back oxidation of CL at C-19, first forming 19-hydroxy-CL and then CLA (Abe et al. 2014). CLA has been reported to accumulate in Arabidopsis roots, including those in atd14 and max2 mutants (Abe et al. 2014). Endogenous CLA has also been reported in rice plants, and exogenous CLA is converted into SLs using the d10-2 rice mutant (Abe et al. 2014). When provided with 13C-labeled CLA, d10-2 mutant root exudates subsequently accumulated 13C-labeled 5-deoxystrigol and orobanchol (Abe et al. 2014). In Arabidopsis, CLA is similarly converted into 5-deoxystrigol and 4-deoxyorobanchol (4DO) (Abe et al. 2014). 5-deoxystrigol is the simplest SL as it lacks hydroxyl, acetyloxyl and other oxygen-containing substituents (Awad et al. 2006; Yoneyama et al. 2008). It is found in both monocots (Awad et al. 2006) and dicots (Yoneyama et al. 2008), indicating it as the precursor of all SLs. 5-deoxystrigol then undergoes either allylic hydroxylation (to strigol or orobanchol) or homoallylic hydroxylation (to sorgomol) (Rani et al. 2008; Xie et al. 2010). Further modification of sorgomol—oxidation of its hydroxymethyl group followed by decarboxylation—results in the formation of sorgolactone (Xie et al. 2010). CLA can also undergo methylation (through an unknown methyl transferase enzyme) and be converted into the methyl ester MeCLA (SL-LIKE1) (Seto et al. 2014). Interestingly, the conversion of CLA into MeCLA is MAX1-independent as confirmed by Arabidopsis mutant analyses (Abe et al. 2014). Another enzyme LBO (Lateral Branching Oxidoreductase) acts downstream of MAX1 to convert MeCLA into the recently identified hydroxymethyl carlactonoate involved in shoot branching (Brewer et al. 2016; Yoneyama et al. 2020).

Recently, a carotenoid-derived molecule zaxinone has been shown to negatively regulate SL (4-deoxyorobanchol) biosynthesis in rice under phosphate (Pi) limiting conditions (Wang et al. 2019). This was confirmed by increased SL content in zaxinone synthase (zas) mutant seedlings under Pi stress and enhanced Striga germination stimulation potential of zas root exudates (Wang et al. 2019). This was similarly observed in tomato root exudates under Pi-deficient conditions (López-Ráez et al. 2008). Enhanced seed germination vigor coincided with increased SL levels, which then decreased upon phosphate restoration (López-Ráez et al. 2008).

Strigolactone Signaling Cascade: A Tale of Binding, Derepression, and Hydrolysis

Phytohormone perception relies on a well-defined receptor system. Just like jasmonate, auxin, and gibberellin signaling (Schwechheimer and Willige. 2009; Dharmasiri et al. 2005; Katsir et al. 2008), SL signaling involves polyubiquitination and proteasomal degradation. The SL signaling cascade involves three important components: (1) an α/β fold hydrolase called D14 in rice (Arite et al. 2009), (2) an F-box leucine-rich protein called MAX2/D3 (Stirnberg et al. 2002; Johnson et al. 2006), and (3) a repressor protein called D53 belonging to the SMAX1-like (SMXL) protein family (Jiang et al. 2013; Stanga et al. 2013). The SL receptor protein D14 is activated after ligand binding, leading to its interaction with other molecules to form a signaling complex; hormonal signal transduction is followed by subsequent hydrolysis of the bound SL, deactivating the hormone (Marzec et al. 2016).

Various SL-insensitive mutants were analyzed to identify different SL signaling components (Seto et al. 2014). AtD14/D14/DAD2 are the orthologous SL receptors in A. thaliana, Oryza sativa, and Petunia, respectively (Waters et al. 2012b; Arite et al. 2009; Hamiaux et al. 2012); gene mutations result in a SL-specific phenotype that is not reversed by GR24 treatment (Arite et al. 2009). These gene orthologs encode proteins similar to the soluble gibberellic acid (GA) receptor GID1 (GIBBERELLIN-INSENSITIVE DWARF1) (Ueguchi-Tanaka et al. 2005). These receptor proteins have a conserved catalytic triad consisting of Ser, His, and Asp (Zhao et al. 2013). GR24 undergoes hydrolysis, most probably due to catalytic triad activity (Kagiyama et al. 2013). The Petunia receptor DAD2 loses its catalytic activity with a Ser-to-Ala substitution (DAD2:S96A) in the triad (Hamiaux et al. 2012), leading to loss of receptor interaction with the F-box protein, thereby suppressing shoot branching (Hamiaux et al. 2012; Marzec et al. 2016). GR24 undergoes very slow hydrolysis with DAD2, but the dad2 mutant phenotype is not reversed by the resulting products (Zhao et al. 2013). This confirms DAD2 involvement in SL signaling, with the hydrolytic process being more important than the end products (Seto and Yamaguchi 2014).

In rice, the SL hormone-D14 receptor interaction results in SL cleavage and subsequent production of a “covalently linked intermediate molecule” (CLIM) bound to D14 (Bythell-Douglas et al. 2017). Unlike other phytohormones, SL signaling depends upon hormone degradation. In detail, binding of D14 with SL leads to nucleophilic attack, resulting in SL ligand dissociation into two molecules: (1) the ABC ring portion called ABC-formyltricycliclactone (ABC-FTL) and (2) the remaining part with the D ring called hydroxymethylbutenolide (HMB) (Nakamura et al. 2013). ABC-FTL is released while HMB remains covalently attached to the D14 receptor; this HMB-D14 intermediate is called CLIM (Yao et al. 2016). This reaction changes the D14 conformation, allowing it to interact with downstream signaling components (Marzec et al. 2019).

SL signaling proceeds from the interaction between the receptor D14 and F-box leucine-rich protein MAX2/D3/RMS4 (orthologs in A. thaliana, Oryza sativa, and Petunia, respectively) (Hamiaux et al. 2012). MAX2 forms a part of the Skp–Cullin–F-box containing (SCF) E3 ubiquitin ligase complex (Hamiaux et al. 2012; Zheng et al. 2014; Zhao et al. 2014). Mutations in these orthologs lead to SL insensitivity, confirming their crucial role in SL signaling (Marzec et al. 2016).

This SCF complex targets the D53 and D53-like SMXL repressor proteins for proteasomal degradation (Jiang et al. 2013; Zhou et al. 2013; Bennett et al. 2016). In Arabidopsis, SMXL6-8 have been proposed to be D53 orthologs, as they regulate shoot branching and other SL-controlled processes (Soundappan et al. 2015; Bennett et al. 2016; Ligerot et al. 2017). Due to its EAR motifs, D53 is expected to interact with TOPLESS-related (TPR) transcriptional corepressor proteins (Smith and Li. 2014). This D53-TPR complex may then repress SL target gene expression (Smith and Li. 2014). The D53 repressor also interacts with the D14 receptor; upon GR24 treatment, D53 undergoes SCF complex-directed degradation (Smith and Li. 2014). The ligand-induced conformational change in D14 allows the receptor to recruit SMXL7 into the SCF complex (Liang et al. 2016). SMXL7 functions both transcriptionally and non-transcriptionally, but the molecular events after its degradation have not been clearly elucidated (Waters et al. 2017; Bythell-Douglas et al. 2017). In O. sativa, the major regulator of plant architecture Ideal Plant Architecture 1 (IPA1) acts downstream of the D53 repressor, regulating SL-induced gene expression (Song et al. 2017). IPA1 is repressed by D53 in vitro and in vivo, which represses its transcriptional activation function (Song et al. 2017).

Several engrossing hypotheses have been proposed to explain the evolution of ligand and signaling specificity by D14 and D14-like receptor proteins. In parasitic plants, D14-like proteins—closely related to D14 proteins—act as receptors of host-exuded SLs, representing a case of convergent evolution (Tsuchiya et al. 2015; Conn and Nelson 2015). These subfamilies of D14-like proteins also include subfunctionalized proteins that respond to other ligands, such as karrikins and other d-lactone-containing compounds (Waters et al. 2012b; Saeed et al. 2017). Perception of both SLs and karrikins also requires the MAX2 F-box protein (Zhao et al. 2015). However, it is unknown how MAX2 discriminates between the two pathways to generate different responses, because F-box proteins tend to be indiscriminate when recruiting target proteins (Nelson et al. 2011; Nakamura et al. 2013). Wang et al. (2020b) proposed that in Arabidopsis, both SL and karrikin signaling pathways converge at SMXL2, as it acts as their common target for polyubiquitination and degradation in a D14- or KAI2-dependent manner.

Different lines of evidence support the model that SL signal transduction occurs as a result of SL binding/hydrolysis-induced conformational changes in the D14 receptor. For example, thermal destabilization of the D14 receptor is initiated by GR24, which depends on an intact D14 catalytic triad (Waters et al. 2015). GR24 also promotes the physical interaction between MAX2/D3 and D14, with MAX2/D3 further destabilizing the D14 receptor (Waters et al. 2017; Zhao et al. 2014). Interestingly, D14-D3 association in O. sativa is a bit more responsive to 2ʹR stereoisomers of SL analogs compared to 2ʹS stereoisomers (Zhao et al. 2015). Furthermore, there are no major structural differences between D14 and apo-D14, when associated with 5-hydroxy-3-methylbutenolide, 2, 4, 4, trihydroxy-3-methyl-3-butenal or SL (Nakamura et al. 2013).

Recently, several modes of SL–D14 interaction have been determined, but it is unclear how D14 functions with D3 in ubiquitinating the D53 repressor. D3 has a C-terminal α-helix that exists in either engaged or dislodged forms (Shabek et al. 2018). The engaged form enables D14 and D3 binding with a hydrolyzed SL intermediate, while the dislodged form recognizes unmodified D14 and prevents its enzymatic activity (Shabek et al. 2018). The D3 α-helix helps D14 in recruiting D53 in a SL-dependent manner, which then activates the hydrolase (Shabek et al. 2018). The self-induced D14 degradation by SLs (through MAX2) limits their own signaling through a negative feedback loop (Chevalier et al. 2014; Koltai 2014).

Controversially, this CLIM model has been challenged by various experimental evidence. CLIM cannot be accommodated in the D14 active site due to its very small electron density; instead, iodine (I) in the crystallization reagents is suspected to bind the active site (Carlsson et al. 2018). D14-mediated SL hydrolysis is also too sluggish after SL treatment, in sharp contrast to the rapid degradation of target proteins (D53/SMXLs) (Seto et al. 2019). Therefore, the rapid response of SLs cannot be entirely explained by this CLIM model. Instead, it has been recently reported that binding of a complete SL molecule, not a hydrolyzed one, initiates the active D14 receptor signaling; D14 then hydrolyzes SL molecules only after completing the pathway (Seto et al. 2019). Kinetic analysis of the AtD14-catalyzed hydrolysis of 5-deoxystrigol detected two hydrolytic products, ABC-FTL and HMB, as described earlier (Hamiaux et al. 2012). The Kcat, Km and Vmax values were found to be 0.12 min−1, 4.9 μM, and 4.0 nmol/min/mg protein, respectively (Seto et al., 2019). In addition, 3,6′-dihydroGR24, which has a single bond instead of a double bond in the enol ether bridge, is not hydrolyzed by the SL receptors in rice and Arabidopsis (Umehara et al. 2015). Furthermore, D14 catalytic activity is quite low for debranones (SL analogs without the enol ether bridge), but these analogs interestingly yield the same results as GR24 (Scaffidi et al. 2014). These observations raise questions about the role of hydrolysis (by D14) in SL signaling.

Therefore, D14 has a dual function and a new mechanism of SL signaling perception could be proposed (Yao et al. 2016). In molecular detail, the D14 conformational change enlarges the catalytic pocket, allowing SL movement into this pocket and then closing the helical lid domain (Shabek et al. 2018). When a SL molecule binds to the D14 receptor protein, D14 initially attains an unstable conformation due to interruption in the catalytic triad formation (Yao et al. 2016). In this changed conformation, the D14 receptor interacts with other components to carry out the SL signaling cascade (Fig. 2). After activation, D14 (through the surface of its rearranged lid domain) interacts with the F-box protein MAX2/D3 and then D53/SMXL repressor binding occurs around the region of the Asp loop (Seto et al. 2019). After D53/SMXL degradation, the D14 catalytic triad is again reconstructed, which performs the important hydrolysis step, resulting in SL deactivation (Seto et al. 2019). This hormonal degradation mechanism is also found in other hormonal pathways (like GA) and is very important for hormone homeostasis (Yamaguchi. 2008).

Fig. 2
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The SL biosynthetic pathway showing key enzymes and intermediates

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The evolution of the SL signaling mechanism provides informative insights. It is believed that the initial role of SLs was AM fungal recruitment to facilitate more efficient nutrient uptake; this symbiotic association was present in land plants about 360–450 million years ago (Waldie et al. 2014; Simon et al. 1993). Remarkably, SLs are found in algae and SL application results in rhizoid elongation—a response also reported in liverworts and mosses belonging to bryophytes (Delaux et al. 2012); however, it is most probably independent of MAX2 (Waldie et al. 2014). In higher plants, MAX2-independent SL signaling has also been reported. Minute GR24 concentrations can inhibit root growth in the max2 mutant (Shinohara et al. 2013). In charophytes, a D14 member is more closely related to the KARRIKIN INSENSITIVE2 (KAI2) receptor than to canonical D14 proteins (Waldie et al. 2014; Waters et al. 2012b). It might be possible that SLs use this receptor instead of MAX2 to initiate their response (Waldie et al. 2014). The D14 and MAX2 gene clades arose quickly when land plants emerged, with D14 probably appearing due to duplication in the clade, while another duplication within D14 resulted in the evolution of the D14-LIKE2 group (Waters et al. 2012b; Waldie et al. 2014). These duplication events correlate with varying functions as land plants diversified. D53 protein evolution also follows a similar pattern. The D53-like genes in mosses have higher similarity to SMAX1 than to D53/SMAXL7 clade; these clades were then subjected to further duplications (Zhou et al. 2013). Intriguingly, the entry of MAX2 into the SL pathway has not been fully elucidated. It is postulated that MAX2 was initially involved in AM colonization only and its role in SL signaling evolved later (Challis et al. 2013); this is supported by the d3 rice mutant which cannot be colonized by AM fungi (Waldie et al. 2014).

Strigolactone Receptors: Highly Conserved in Diverse Plant Species

The SL receptors have a conserved α/β hydrolase functional domain (Bennett and Leyser 2014), which was first identified in the SL-insensitive O. sativa d14 mutant (Arite et al. 2009). Orthologs were eventually identified in Petunia (Hamiaux et al. 2012), pea (de Saint Germain et al. 2016) and Arabidopsis (Waters et al. 2012b). According to Arite et al. (2009), D14 homologs are found in diverse plant clades, such as Marchantia polymorpha (bryophytes), Selaginella moellendorfii (pteridophytes), and gymnosperms. These homologs belong to the D14-like subfamily, whereas angiosperm genes are grouped into the D14 subfamily of the α/β-hydrolase superfamily (Arite et al. 2009). Proteins of these subfamilies similarly possess a conserved catalytic triad, a nucleophilic residue and an acidic residue, but have quite different sequences (Nardini and Dijkstra. 1999; Arite et al. 2009). The α/β hydrolase superfamily also includes the acetylcholinesterase (AChE) enzyme (responsible for acetylcholine metabolism) and the inactive gibberellic acid receptor (Holmquist et al. 2000).

D14 (without any prefix corresponds to the O. sativa receptor) acts as a receptor as well as an enzyme, differentiating it from other plant hormone receptors (Hamiaux et al. 2012). It has a α/β hydrolase functional domain containing the Ser-His-Asp catalytic triad, forming its ligand binding pocket, and 4 α helices forming its cap (Kagiyama et al. 2013). It consists of 318 amino acids, and a homolog called D14-like is also reported in the rice genome (Arite et al. 2009. The rate of SL hydrolysis in vitro is as low as ~ 0.3 molecules per minute, suggesting that bioactive SL-derived signal production is not its primary function (Snowden and Janssen. 2016). Consistent with this, neither the intermediate molecule 2,4,4-trihydroxy-3-methyl-3-butenal nor the end products of SL hydrolysis (tricyclic lactone and HMB) act as signals for shoot branching suppression (Waters et al. 2017).

The SL receptor in A. thaliana (AtD14) is evolutionarily conserved (Waters et al. 2012b; Arite et al. 2009); just like the rice D14 receptor, it consists of a catalytic triad and possesses both receptor and enzyme functions (Hamiaux et al. 2012). The structure of the AtD14-D3-ASK1 complex showed a portion of the hormone covalently bonded with the receptor through two amino acids in the triad (Yao et al. 2016). When the receptor conformation changes, an α helix domain increases in length, while another α helix domain unfolds and forms a loop (Yao et al. 2016). Four α helix domains form the lid of the receptor, which probably functions in destabilizing the SL receptor upon hormone attachment (Zhao et al. 2015; Snowden and Janssen 2016). The enzymatic active site also decreases in volume resulting in closure (Fig. 3). Therefore, this indicates that D-ring separation is difficult without complex dissociation and could explain the sluggish enzyme activity (Snowden and Janssen 2016). In Arabidopsis, the AtD14L/KAI2 protein is 51% identical and 75.9% similar to AtD14, but is instead involved in karrikin signaling; unsurprisingly, AtD14L and AtD14 belong to different phylogenetic clades (Waters et al. 2012b).

Fig. 3
figure 3

The SL signaling mechanism showing receptor complex formation and protein modifications

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The Petunia D14 receptor ortholog is DAD2 (Simons et al. 2007). Hamiaux et al. (2012) solved its structure by X-ray crystallography and its lid consists of 4 α helices, connected by a β hairpin to the core. A strongly hydrophobic cavity between the lid and the core can easily accommodate known SLs (Hamiaux et al. 2012). The authors further reported that when GR24 is present, DAD2 interacts with the F-box protein PhMAX2A (the Petunia MAX2 ortholog). GR24 then undergoes hydrolysis upon DAD2 interaction, but mutations in the catalytic triad lead to loss of enzymatic activity and failure to interact with PhMAX2A (Hamiaux et al. 2012). The prolific branching phenotype of dad2 mutants has also been observed in dad1 (CCD8) and dad3 (CCD7) biosynthetic mutants (Napoli et al. 1996). DAD2 locally controls shoot branching, as confirmed by grafting and genetic studies (Simons et al. 2007; Hamiaux et al. 2012). The branching phenotype of biosynthetic mutants is reversed by grafting with wild-type root stocks; however, this reversion does not occur in dad2 mutants, suggesting that DAD2 is not involved in SL biosynthesis (Simons et al. 2007).

The SL receptor in Hordeum vulgare (barley) is encoded by the HvD14 gene, which consists of a 1055-bp coding sequence with two exons (Marzec et al. 2016). The approximately 303-amino acid HvD14 protein also contains the conserved α/β-hydrolase domain between amino acids 57 and 295 (Kagiyama et al. 2013). Unsurprisingly, it has great structural similarity, high sequence conservation, and comparable secondary domains to the rice D14 ortholog ((Marzec et al. 2016). In hvd14.d mutants, the Gly at position 193 is substituted by Glu (Marzec et al. 2015); this residue is present in the αD2 α-helical domain, which constitutes the cap surrounding the active site along with αD1, αD3, and αD4 (Kagiyama et al. 2013).

Zheng et al. (2016) reported that the woody perennial plant Populus trichocarpa has two highly identical (91.7%) and similar (95.9%) homologs PtD14a and PtD14b. They showed that PtD14a is 79% identical and 89.1% similar to AtD14, while PtD14b is 77.5% identical and 89.1% similar to AtD14 (Zheng et al. 2016). The crucial Ser-His-Asp catalytic triad is conserved in both PtD14 homologs at positions 96, 246, and 217 (Zheng et al. 2016). In terms of gene expression, PtD14a transcript levels are higher compared to PtD14b, with very low co-expression between them (Zheng et al. 2016).

The probable SL receptors in parasitic weeds were more difficult to identify, because the phenotypes could not be dissected genetically (Toh et al. 2015; Tsuchiya et al. 2015). Subsequently, a group of α/β-hydrolases ShKAI2s/ShHTLs (S. hermonthica KARRIKIN INSENSITIVE2/ HYPO-SENSITIVE TO LIGHT) were discovered to be involved in SL hydrolysis and SL-induced seed germination; these hydrolases are D14 paralogs that act as SL receptors (Conn et al. 2015; Toh et al. 2015; Yao et al. 2017). Among them, ShHTL7 serves as the most active SL receptor in Striga (Conn et al. 2015; Yao et al. 2017). During CLIM formation, ShHTL7 undergoes a conformational change (like AtD14) to transduce signaling through its interaction with MAX2/ShMAX2 (Yao et al. 2017).

Strigolactone-Phytohormone Crosstalk: Dynamic Interplay for Effective Plant Physiology

Different hormonal signaling pathways interact with one another, affecting their respective signaling components (Huot et al. 2014). These dynamic interactions regulate hormonal biosynthesis, response, and transport, thereby helping plants control their morphology and adapt to changing environmental conditions (Cheng et al. 2013). These challenging conditions include severe nutritional deficiency, abiotic stress factors (i.e., salinity, heat, cold, drought, and light stress), and harmful biotic invasions (i.e., pathogens and pests). Phytohormone crosstalk facilitates appropriate and tunable plant responses to these conditions by controlling nutrient distribution and by modulating growth, developmental, and defense processes. Plant stress responses are primarily regulated by jasmonic acid (JA), ABA, and salicylic acid (SA), whereas plant growth/developmental processes are mainly governed by auxins, gibberellins and cytokinins (Huot et al. 2014). SLs interact with other hormones in order to exert their impact (Saeed et al. 2017; Torres-Vera et al. 2014).

Strigolactones and Auxins

SLs inhibit shoot branching by regulating auxin transport. Compared to wild-type plants, A. thaliana max mutants show increased auxin transport due to increased PIN1/3/4/6 gene transcription (Bennett et al. 2006; Lin et al. 2009). Treating Arabidopsis max mutants and rice dwarf mutants with an auxin transport inhibitor, N-1-naphthylphtalamic acid, causes inhibition of bud outgrowth (Cheng et al. 2013; Lin et al. 2009). Crawford et al. (2010) reported that treatment with basal GR24 levels reduces auxin transport basipetally, as well as PIN1 accumulation in xylem parenchyma cell membranes. These observations persist in biosynthetic max1 mutants but not signaling max2 mutants, indicating that SLs slow down polar auxin transport stream in a MAX2-dependent manner (Crawford et al. 2010).

Studies of auxin and max mutants showed that SLs directly affect secondary growth activity, independent of auxin stacking (Agusti et al. 2011), by affecting interfascicular cambium activity (Ruyter-Spira et al. 2011). Based on a quantitative study, max mutants have a 30% decrease in interfascicular cambium-derived tissues, concomitant with lower expression levels of cambium- and cell cycle-related genes (Agusti et al. 2011). SLs regulate auxin content in the primary root tip, because the primary root lengths of SL biosynthetic and signaling mutants are shorter compared to wild-type plants (Ruyter-Spira et al. 2011). GR24 application rescues this short root phenotype in SL-deficient mutants, but not in SL-insensitive max2 mutants (Ruyter-Spira et al. 2011). SLs inhibit auxin efflux by controlling PIN activity, leading to auxin accumulation inside the primary root meristem cells and ultimately resulting in increased primary root length (Ruyter-Spira et al. 2011). SL–auxin interaction controls root development by adjusting or regulating intercellular auxin flow, auxin sensitivity, and shoot-to-root transport (Mayzlish-Gati et al. 2012; Omoarelojie et al. 2019). SLs also control lateral root formation by adjusting the essential auxin gradient (Omoarelojie et al. 2019). Furthermore, SL–auxin interaction regulates root hair elongation, whereby SLs increase intracellular auxin concentration by hindering auxin efflux (Koltai et al. 2010). Ligerot et al. (2017) suggested that a feedback loop exists in the auxin-SL crosstalk. Auxins upregulate SL biosynthesis in an RMS2 (encodes PsAFB4/5 auxin receptor)-dependent manner, while SLs downregulate auxin levels in an RMS3- and RMS4-dependent manner by downregulating auxin biosynthetic gene expression (Ligerot et al. 2017).

Pi deficiency leads to increased levels of RSL4, an auxin-related transcription factor that promotes root hair elongation (Omoarelojie et al. 2019; Datta et al. 2015). In contrast to auxins, SLs inhibit adventitious root (AR) formation in Arabidopsis and pea (Datta et al. 2015). AR inhibition was even evident with high auxin concentration, suggesting that suppression of AR formation is not due to low auxin levels (Rasmussen et al. 2012). Auxins and SLs also play a crucial role during mycorrhization; auxins are associated with arbuscule formation, whereas SLs are associated with presymbiotic fungal growth (Guillotin et al. 2017). The authors further found that auxin content increases in roots colonized by AM fungi, and exogenous auxin application promotes the colonization process. An auxin-related gene Sl-IAA27 positively controls mycorrhization by regulating SL biosynthesis via NSPI (transcription factor of the D27 and MAX1 genes) (Guillotin et al. 2017).

Strigolactones and Cytokinins

Cytokinins are adenine-derived plant hormones that stimulate cytokinesis and influence various processes, like enhancing shoot growth, limiting root growth, and influencing axillary shoot branching (Aloni et al. 2006; Werner et al. 2001). In P. sativum and A. thaliana, branching mutants with increased SLs have reduced cytokinin concentrations in the xylem sap (Morris et al. 2001; Foo et al. 2007). Decreased cytokinin sensitivity has also been reported in the buds of SL-insensitive plants (El-Showk et al. 2013). Dun et al. (2012) reported that the SL-insensitive and SL-deficient P. sativum rms mutants (rms4 and rms1) have increased expression of the cytokinin biosynthetic gene PsIPT1 in shoot nodes and internodes. Interestingly, the rms1 mutant was more sensitive to low cytokinin levels compared to wild type, when applied to the buds or supplied through the vasculature (Dun et al. 2012). The authors further found that bud outgrowth is higher in rms1 mutants than wild-type plants after applying low cytokinin levels, suggesting that SLs and cytokinins play antagonistic roles. Exogenous GR24/ cytokinin application weakened the effect of cytokinins in rms1 mutants but not in rms4 mutants, implying that SL-cytokinin interaction converges at RAMOSUS4 (RMS4) (Dun et al. 2012). The cytokinin-SL antagonism is due to PsBRC1, a common target of both hormones (El-Showk et al. 2013); its gene expression negatively correlates with bud growth (Dun et al. 2012). Additionally, PsBRC1 gene expression is enhanced by GR24 but reduced by cytokinins – a trend that persists even with cycloheximide (ribosomal translation inhibitor) treatment, suggesting that new protein synthesis is not required for this regulation (Dun et al. 2012). Both SLs and cytokinins act as negative regulators of lateral root development; the cytokinin receptors ARR1, ARR12, and AHK3 are associated with GR24-induced reduction of lateral development (Ruyter-Spira et al. 2011; Jiang et al. 2015). Genetic studies show that GR24-regulated lateral development is influenced by PIN1- and PIN7-mediated auxin polar transport; cytokinin treatment downregulates PIN1/PIN3/PIN5 but upregulates PIN7 expression (Jiang et al. 2015). Moreover, the A. thaliana max2 mutants show low cytokinin catabolic gene expression (CKX1, 2, 3, 5), reflecting the negative relationship between cytokinins and SLs (Banerjee et al. 2018). In O. sativa, Duan et al. (2019) observed enhanced cytokinin levels in shoot bases of d53 mutants.

Some evidence suggests that SLs and cytokinins play important roles during drought adaptation (Nishiyama et al. 2011). Analyses of cytokinin-depleted Arabidopsis mutants (CKX- overexpressor), as well as signaling mutants (arr1, 10, 12), indicated that cytokinin signaling negatively regulates drought acclimation (Nguyen et al. 2016). Drought tolerance mechanisms in these mutants involve amplified stomatal closure, increased root-to-shoot ratio, enhanced cell membrane integrity, and increased ABA hypersensitivity (Nishiyama et al. 2011). Due to the undesirable role of cytokinins in drought tolerance, cytokinin biosynthesis and signaling in A. thaliana are suppressed during drought (Cortleven et al. 2019). Drought-induced cytokinin suppression occurs through the ABA-induced transcription factor AtMYB2, and members of the ABA-activated sucrose nonfermenting 1 (SNF1)-related protein kinase 2 family (Cortleven et al. 2019). In contrast to cytokinins, SLs positively regulate resilience to water stress conditions, as shown in studies of Arabidopsis max1 mutants and CCD7-silenced tomato mutants (Visentin et al. 2016; Zhang et al. 2014). Additionally, SLs decrease stomatal density (Van Ha et al. 2014) and stomatal opening during drought (Zhang et al. 2018). The max mutants also show decreased response to ABA (Van Ha et al. 2014). Overall, these observations clearly indicate the contrasting roles of SLs and cytokinins under drought stress conditions (Li et al. 2019).

Strigolactones and Gibberellins

The phytohormones SLs and gibberellins (GAs) may interact during their perception and signaling, acting together during plant growth and development (Marzec 2017). Remarkably, SL biosynthesis can be regulated by GAs (Ito et al. 2017). GAs are involved in flowering, seed production, leaf morphology, and shoot/root growth (Claeys et al. 2014). Various studies have indicated that SL and GA signaling are very similar. Rice semi-dwarf mutants in GIBBERELLIN OXIDASE 5, 6 and 9 exhibit an extra-branched shoot phenotype similar to SL mutants (Marzec 2017). GAs control tiller number through the action of ORYZA SATIVA HOMEOBOX1 (osHB1) and TEOSINTE BRANCHED1 (osTB1) transcription factors (Lo et al. 2008). SLs promote the interaction between the D14 receptor and SLENDER1 (SLR1), a negative regulator of GA signaling (Nakamura et al. 2013). SLR1 degradation occurs in an SL-dependent manner, which parallels the GA signaling pathway, where the GID1 receptor binds GA to promote interaction between GID1 and DELLA proteins, eventually leading to DELLA degradation via the 26S proteasome (Marzec. 2017). Additionally, gene expression databases show that GA3 treatment decreases SL biosynthetic gene expression in O. sativa (Ito et al. 2017). The interaction between SLs and GAs in A. thaliana is inconclusive; microarray data showed varying SL biosynthetic gene expression profiles upon GA3 treatment (Marzec et al. 2015). In O. sativa Zou et al. (2019), found that SL biosynthetic and signaling mutants exhibit dwarfism that is rescued by GA treatment. Interestingly, these mutants have less bioactive GA and decreased GA sensitivity (Zou et al. 2019). This ultimately leads to reduced shoot length by downregulating genes involved in cell division and elongation (Zou et al. 2019).

Strigolactones and Abscisic Acid

ABA is regarded as a universal stress hormone since it regulates various abiotic stress responses. Like ABA, SLs are apocarotenoid hormones so it is possible that they could also act as stress hormones. Tomato ABA mutants have low SL biosynthetic gene expression, including LeCCD7 and LeCCD8, reflecting the close harmonization between SL and ABA anabolic pathways (Banerjee et al. 2018). SL-deficient Arabidopsis mutants have downregulated ABA import genes, like ABCG22 and ABCG40, resulting in ABA hyposensitivity (Van Ha et al. 2014). It has also been reported that mycorrhizal plants exposed to abiotic stresses have greater SL and ABA levels (Ruiz-Lozano et al. 2016). GR24 application decreased the expression of LjNCED2 in Lotus japonicus, which in turn inhibited ABA accumulation during osmotic stress (Liu et al. 2015). Additionally, SL–ABA interaction is demonstrated by SLs controlling ABA-induced stomatal sensitivity (Van Ha et al. 2014). SLs promote seed germination under high temperature conditions by regulating both ABA and GAs in parasitic and non-parasitic seeds (Mostofa et al. 2018). Furthermore, SL biosynthetic and signaling genes in Sesbania cannabina are upregulated by ABA to cope with salt stress, while SL biosynthetic inhibitor treatment induced partial salt tolerance (Ren et al. 2018). Studies using ABA-deficient tomato mutants and CCD/NCED inhibitors suggest that SL regulates ABA biosynthesis through an unknown mechanism (López-Ráez et al. 2010).

Strigolactones and Ethylene

Certain plant growth and developmental processes involve both SL and ethylene signaling, including seed germination, leaf senescence, root hair elongation, and hypocotyl growth (Ueda and Kusaba 2015; Cheng et al. 2013; Kapulnik et al. 2011). During light treatment, SLs upregulate HY5 expression in a MAX2-dependent fashion, inhibiting hypocotyl elongation (Jia et al. 2014). In contrast, ethylene promotes hypocotyl elongation by augmenting HY5 degradation via COP1 (Yu et al. 2013). These show the antagonistic roles of these two hormones in regulating hypocotyl growth. SL-mediated root hair elongation also depends on ethylene signaling, since ethylene signaling mutants (like At-etr) have reduced GR24 sensitivity (Kapulnik et al. 2011). Abolishing ethylene production totally eliminates SL-mediated root hair elongation, while GR24 enhances ethylene biosynthetic gene ACS2 transcription (Kapulnik et al. 2011). Moreover, SLs stimulate ethylene biosynthesis in Striga seeds prior to germination (Sugimoto et al. 2003). During leaf senescence, SLs activate senescence signals mediated by ethylene (Ueda and Kusaba 2015).

Strigolactones and Salicylic Acid

SA is involved in plant defense responses against various pathogens, as well as tolerance to abiotic stresses (Askari and Ehsanzadeh 2015; Prodhan et al. 2018; Omoarelojie et al. 2019). SA-mediated stress tolerance is mainly due to changes in the plant’s reactive oxygen species status (Omoarelojie et al. 2019). In terms of crosstalk, SA interacts with SLs during plant–fungal symbioses (Rozpadek et al. 2018). GR24 treatment results in SA build-up, whereas max2 mutants have decreased SA concentrations, suggesting that SLs are involved in plant defenses by inducing SA production (Rozpądek et al. 2018; Omoarelojie et al. 2019). In wheat, foliar application of SLs and SA synergistically results in lower electrolyte leakage, higher relative leaf water content, and enhanced antioxidant enzyme activities during drought stress (Sedaghat et al. 2017).

Strigolactones and Jasmonic Acid

Jasmonates are involved in secondary metabolism, wounding responses, and plant–pathogen/insect interactions (Yan et al. 2007; Yan and Xie. 2015). JA concentration and JA-dependent PIN11 gene expression are reduced in the tomato SL biosynthetic mutant Sl-ccd8 (Torres-Vera et al. 2014). Because PIN11 provides resistance in Solanum lycopersicum against Botrytis cinerea (Torres-Vera et al. 2014), these observations hint at a possible interplay between these two hormones during disease resistance. Although there is no direct evidence depicting SL–JA interaction, both are involved together in several processes, like plant–microbe interactions, mesocotyl elongation, and senescence; thus, their crosstalk cannot be totally ruled out (Omoarelojie. et al. 2019). For example, Lahari et al. (2019) reported that SLs induce root-knot nematode infection in rice roots by inhibiting the JA pathway. Remarkably, SL biosynthetic mutants were less prone to infection by the root-knot nematode Meloidogyne graminicola (Lahari et al. 2019).

Strigolactones and Karrikins

Karrikins (from ‘karrik’ meaning smoke) or KARs are smoke-derived signals produced by burning vegetation; they form through the combustion of carbohydrates (Flematti et al. 2011). Although not produced in planta, they can stimulate germination of dormant seeds (De Cuyper et al. 2017) – an effect attributed to the butenolide pyran moiety (Flematti et al. 2007). Unlike SLs, however, KARs do not induce the germination of parasitic weeds (Conn et al. 2015). Although they have different sources and effects on plant growth and development, SLs and KARs share highly similar signaling mechanisms, which could be due to their shared butenolide structure (Morffy et al. 2016). The KAI2 receptor of KARs work in the same manner as the D14 receptor of SLs (Morffy et al. 2016). Because KAI2 and D14 are paralogs, they share the F-box protein MAX2 during signaling (De Cuyper et al. 2017). Structurally, the KAI2 receptor catalytic pocket is smaller than that of the D14 receptor, which hints at the binding of smaller cognate molecules (Guo et al. 2013). Phylogenetic studies have shown that KAI2 was present in basal land plants instead of D14 orthologs, suggesting that KAI2 is ancestral and that D14 probably evolved due to KAI2 duplication (Waters et al. 2012b).

The application of KAR1, KAR2, as well as rac-GR24 inhibits hypocotyl elongation in Arabidopsis, with rac-GR24 having greater impact than KARs (Nelson et al. 2010; De Cuyper et al. 2017). This observation is supported by max2 mutant plants that have longer hypocotyls (Stirnberg et al. 2002), a phenotype shared by mutant kai2 seedlings (Waters et al. 2012b). In contrast, KAR1 and rac-GR24 have antagonistic effects on cotyledon growth – karrikin promotes growth while rac-GR24 negatively impacts cotyledon growth (De Cuyper et al. 2017). Mutations in KAI2 and MAX2 cause skewing of A. thaliana roots, but this response is independent of SL perception by the D14-MAX2 complex (Swarbreck et al. 2019). Scaffidi et al. (2014) cautioned about using racemic mixtures of chemically synthesized SLs, as well as their analogs like GR24, since they can activate responses that are different from natural counterparts.

As reported by Liu et al. (2019), both SLs and KARs shape the morphology of the exodermis. They revealed that SLs positively regulate the number of hypodermal passage cells (HPC), but d14 mutants surprisingly have higher HPCs (Liu et al. 2019). They further noted that, in contrast to d14, max2 mutants have decreased HPC numbers (Liu et al. 2019). In Petunia, KAI2 mutation also reduces HPC numbers, indicating the critical importance of the dimeric KAI2/MAX2 receptor in controlling this process (Liu et al. 2019).

Strigolactones and Nitric Oxide

There is evidence that SLs and nitric oxide (NO) possibly interact during various stress responses and developmental processes. Their interplay has mostly been studied in root systems; results suggest that NO negatively and positively regulates root SL biosynthesis and signaling, respectively, in a nutrient-dependent manner (Bharti and Bhatla. 2015). NO can modify proteins involved in SL biosynthesis and signaling, with Arabidopsis max1-1 and max2-1 mutants having increased NO levels in their root tips (Kolbert. 2019). These observations highlight the possible negative impact of SLs on NO biosynthesis; however, exogenous SL application increased NO production, contradicting earlier genetic studies (Kolbert. 2019). GR24 treatment results in decreased NO concentration in lateral roots but increased NO concentrations in primary root tips (Bharti and Bhatla. 2015). Furthermore, SLs and NO act as positive regulators of meristem activity thereby enhancing root elongation (Sun et al. 2016). Endogenous NO does not influence SL biosynthesis, while exogenous NO upregulates the expression of SL signaling but not biosynthetic genes in O. sativa (Sun et al. 2016). In addition, exogenous SLs promote accumulation of guard cell H2O2 and NO, leading to SLOW ANION CHANNEL-ASSOCIATED 1-mediated stomatal closure (Lv et al. 2017).

Conclusion and Future Prospects

SLs regulate plant growth, development, and stress tolerance via close crosstalk with other hormones. Mechanistically, SLs elicit their response by regulating hormone content, transport, and delivery between diverse plant organs and within plant tissues, and also by interacting with other hormone signaling cascades. Plant responses are governed by synergistic as well as antagonistic interactions of SLs with other phytohormones. Based on various physiological and molecular studies, SLs are essential for plant responses to stressful environmental conditions. Due to their utmost importance, continued research is needed to more lucidly understand the SL biosynthetic pathway, SL signaling crosstalk with other hormones, and mechanisms by which SLs regulate different stress responses, growth processes, and developmental programs. Although we have gained significant insights in understanding SL hormonal interplay at various levels of regulation, critical knowledge gaps still need to be addressed at both cellular and molecular levels. Certain functions of SLs have yet to be discovered, while further investigating the SL repressor D53 could reveal its involvement in other processes. On a translational level, studying SL hormones could help produce crop varieties with better nutrient allocation under limiting conditions. Long-term research programs could focus on developing more resilient crops, through genetic manipulation of SL quantity and response. Moreover, whether the SL receptor enzymatic activity is required for downstream SL signaling and function still needs to be elucidated. Because protein–protein interactions during SL signaling are unique, further research is required to fully understand SL crosstalk with other hormone pathways. To gain better insights and solve pressing biological problems, the next decade opens a lot of research opportunities in the exciting field of strigolactone hormone biology.