Strigolactone Synthesis Essay

Abstract

Strigolactones (SLs), first identified for their role in parasitic and symbiotic interactions in the rhizosphere, constitute the most recently discovered group of plant hormones. They are best known for their role in shoot branching but, more recently, roles for SLs in other aspects of plant development have emerged. In the last five years, insights into the SL biosynthetic pathway have also been revealed and several key components of the SL signaling pathway have been identified. Here, and in the accompanying poster, we summarize our current understanding of the SL pathway and discuss how this pathway regulates plant development.

Introduction

Strigolactones (SLs) are evolutionarily ancient plant signaling molecules that play roles in diverse organisms [such as bryophytes, angiosperms and arbuscular mycorrhizal (AM) fungi] and in several aspects of plant development. The name ‘strigolactone' comes from the first identified role of these compounds as stimulants of seed germination in species of the parasitic weed Striga (Cook et al., 1966) and from their lactone ring-containing chemical structure. In 2008, SLs were shown to play a role in repressing shoot branching and hence were included in the list of plant hormones that modulate plant development. These early studies showed that SL-deficient mutants are highly branched and that application of the synthetic SL GR24 inhibits axillary bud outgrowth (Gomez-Roldan et al., 2008; Umehara et al., 2008). Since then, novel functions of SLs in plant development have continuously been identified (Brewer et al., 2013; Ruyter-Spira et al., 2013). Furthermore, parts of the SL biosynthesis pathway have recently been deciphered, with the discovery of carlactone (CL) as a key intermediate for SL biosynthesis as well as of novel bioactive SL-like compounds (Abe et al., 2014; Al-Babili and Bouwmeester, 2015; Seto et al., 2014). Receptors for SLs, and some of their downstream effectors, have also been identified (Hamiaux et al., 2012; Xiong et al., 2014).

Here, we summarize our current understanding of the SL biosynthesis and signaling pathways, we describe the different functions of SLs during plant development, and we discuss the evolutionary origin of SL signaling. Models of SL signaling and crosstalk with other plant signaling pathways in the context of shoot branching are also presented.

Chemical structure and biosynthesis of natural SLs

The structural core of SLs is a tricyclic lactone (containing rings that are referred to as ABC), with different carbon A-ring sizes and substitution patterns on AB-rings. This core is connected via an enol ether bridge to an invariable α,β-unsaturated furanone moiety (termed the D-ring). To date, at least 20 naturally occurring SLs have been identified and characterized in root exudates of various land plants (Al-Babili and Bouwmeester, 2015; Tokunaga et al., 2015). They can be separated into two types – strigol and orobanchol – according to the stereochemistry of the B–C-ring junction, with both having a conserved R-configuration at the C-2′ position. The bioactiphore responsible for the various SL bioactivities resides within the region that connects the D-ring to the core, which can differ according to SL type (Zwanenburg and Pospíšil, 2013).

Recent studies have provided some insights into the SL biosynthesis pathway. SLs are synthesized from the key precursor CL, which is derived from all-trans β-carotene via the action of an isomerase (D27) and two carotenoid cleavage dioxygenases (CCD7 and CCD8). These steps of the SL biosynthesis pathway take place in the plastid and the resulting CL is then exported into the cytoplasm. The subsequent steps involve CL oxidation, further ring closures and functionalizations involving members of the CYP711 family (MAX1) (Zhang et al., 2014), eventually giving rise to SLs and SL-like compounds. In Arabidopsis, the unique enzyme MAX1 is able to transform CL to carlactonoic acid, which is further transformed into the SL-like compound methyl carlactonoate (MeCLA) by an unknown enzyme (Abe et al., 2014). Other SL-like compounds, with a CL-type structure lacking the canonical ABC-rings (Kim et al., 2014; Ueno et al., 2014), have been discovered in different plants, highlighting the structural diversity of this class of compounds. Once synthesized, all of these compounds may be transported within the plant and in the rhizosphere. PhPDR1, a member of the ABC family, has been identified as a potential SL transporter (Kretzschmar et al., 2012; Sasse et al., 2015).

SL biosynthesis, which occurs mainly in roots but also in the stem, is tightly regulated (Al-Babili and Bouwmeester, 2015). Phosphate starvation, for example, strongly induces SL biosynthesis (López-Ráez et al., 2008). In addition, high levels of SL biosynthesis gene transcripts (in particular CCD7 and CCD8) in SL-deficient and SL-insensitive mutants in several species indicate that there is feedback regulation of SL biosynthesis (Hayward et al., 2009; Proust et al., 2011).

SL perception: coordinating enzymatic activity and reception

Using genetic approaches, genes encoding SL receptors have been identified in several vascular land plants, including petunia (where the receptor is named DAD2), rice (D14) and Arabidopsis (AtD14) (Arite et al., 2009; Hamiaux et al., 2012; Waters et al., 2012). These proteins belong to a clade of the α/β-hydrolase enzyme superfamily that includes DWARF14 (D14) and GID1, which is a gibberellin receptor that has lost its enzymatic activity. It was shown, using in vitro enzymatic studies, that D14 proteins can hydrolyze the synthetic SL GR24 into inactive ABC- and D-ring parts (Hamiaux et al., 2012; Seto and Yamaguchi, 2014; Xiong et al., 2014). Importantly, it was shown that the SL-like molecule MeCLA can also interact in vitro with the AtD14 SL receptor and be hydrolyzed, despite not having the canonical four-ring structure (Abe et al., 2014). The enzymatic activity of D14 proteins is mediated by a conserved catalytic triad, Ser-His-Asp (S-H-D), that has hydrolase activity. Additionally, the catalytic triad seems to be important for the biological response of D14 proteins since mutated proteins, at least those harboring mutations at the Ser residue, cannot complement the d14 mutant branching phenotype, as shown in petunia (Hamiaux et al., 2012).

The mechanism of SL reception by D14 is still not well understood; in particular, it is unclear whether SL hydrolysis by the receptor is of importance as the hydrolysis products have been shown to be biologically inactive. However, it has been proposed that, following hydrolysis, the D-OH part of SLs forms a complex with D14 thereby allowing the recruitment of binding partners (de Saint Germain et al., 2013; Nakamura et al., 2013).

It should be noted that the D14 SL receptor clade is closely related to the KARRIKIN INSENSITIVE 2/HYPOSENSITIVE TO LIGHT (KAI2/HTL) clade. In Arabidopsis, KAI2 is able to perceive butenolide-containing rings, including the smoke-derived karrikin (KAR) compounds, which, similar to SLs, have a lactone D-ring (Guo et al., 2013b; Smith and Li, 2014; Waters et al., 2012). Interestingly, the SL and KAR pathways modulate plant development in distinct ways but both require the F-box protein MAX2 (D3) to mediate their responses (Nelson et al., 2011).

SL signaling: the role of UPS-dependent protein degradation

Most plant hormone signaling pathways involve the targeting of proteins for degradation through the ubiquitin-proteasome system (UPS) (Kelley and Estelle, 2012). There are strong arguments to suggest that the UPS is also involved in SL signaling (Bennett and Leyser, 2014). In particular, the F-box protein MAX2, which is part of a SKP1–CULLIN–F-BOX (SCF) ubiquitin ligase protein complex, appears to play a key role in mediating SL-triggered protein degradation (Stirnberg et al., 2007; Zhao et al., 2014). Recently, the protein D53 was identified in rice and shown to be targeted for degradation after SL treatment; this degradation was not observed when the proteasome inhibitor MG132 was used (Jiang et al., 2013; Zhou et al., 2013). The rice d53 semi-dominant mutant, which expresses a mutated protein that is resistant to degradation by SL treatment, is SL insensitive and shows high tillering/branching (Jiang et al., 2013; Zhou et al., 2013). Furthermore, it was shown that a reduction in D53 expression in d3 and d14 mutant backgrounds can restore a non-branched wild-type phenotype (Jiang et al., 2013; Zhou et al., 2013). Together, these data indicate that D53 acts as a suppressor of SL signaling in the control of shoot branching (Jiang et al., 2013; Zhou et al., 2013).

This idea led to a model in which MAX2 interacts with D14 in an SL-dependent manner, and this leads to the ubiquitylation-dependent degradation of D53 by the SCFMAX2 complex. However, nothing is currently known about the subsequent effects of D53 degradation and how this protein acts to suppress SL signaling. D53 belongs to a small family of proteins [SMAX1-like (SMXL)] that show some homology with class I CIp ATPase enzymes (Stanga et al., 2013). The presence of potential ethylene-responsive element binding factor-associated amphiphilic repression (EAR) motifs in D53 and its ability to interact with topless-related (TPR) proteins, which are known transcriptional co-repressors, suggest that a D53-TPR complex could repress the transcription of downstream targets (Bennett and Leyser, 2014; Jiang et al., 2013; Smith and Li, 2014), but this has not yet been demonstrated. Other proteins that are subject to SL-triggered degradation, or other transcription factors that lie directly downstream of D53/SMXL proteins, are still unknown (Smith and Li, 2014). BES1, a positive regulator in the brassinosteroid signaling pathway, can also be targeted for degradation via SCFMAX2, although SL is not needed for a BES1-MAX2 interaction (Wang et al., 2013). An SL-dependent interaction between SLR1, a rice gibberellin signaling repressor (DELLA protein), and D14 has also been shown but the biological significance of this interaction is not yet understood (Nakamura et al., 2013).

Models for SL signaling in the control of shoot branching

SLs are best known for their role in repressing shoot branching, and two mechanisms have been proposed to explain this role. In rice and pea, SLs were shown to act via their effects on the TCP transcription factor OsTB1/PsBRC1 to repress axillary bud outgrowth (Braun et al., 2012; Minakuchi et al., 2010). This transcription factor acts as a key integrator of several other pathways, such as the cytokinin pathway and the recently proposed sucrose signaling pathway in pea (Mason et al., 2014; Rameau et al., 2015). Interestingly, the maize ortholog (TB1) of the gene encoding this transcription factor seems to act independently of SLs to repress shoot branching (Guan et al., 2012). In rice, other transcription factors, such as MADS57 and IPA1/OsSPL14, that are involved in shoot branching have also been connected to key components of the SL signaling pathway, but whether these various transcription factors mediate SL signaling, and if or how they lie downstream of the D14-D3-D53 axis, are still not clear (Guo et al., 2013a; Lu et al., 2013).

Because SL-deficient mutants are more branched than brc1 mutants (Braun et al., 2012; Minakuchi et al., 2010), there is very likely a BRC1-independent effect of SLs on shoot branching. Indeed, in Arabidopsis a non-transcriptional mechanism relies on SLs triggering the rapid removal of the auxin efflux carrier PIN-FORMED 1 (PIN1) from the plasma membrane of stem xylem parenchyma cells (Shinohara et al., 2013). This effect of SLs would increase competition between buds to export auxin into the main auxin stream, based on an auxin transport canalization-dependent mechanism (Shinohara et al., 2013; Waldie et al., 2014).

Whether both of these mechanisms regulate shoot branching or act at different stages of bud outgrowth is still debated. Moreover, whether these downstream targets of SL signaling are dependent on the UPS-mediated degradation of D53 remains to be clarified.

Key developmental roles for SL signaling

SLs control numerous other aspects of plant development. Pea, Arabidopsis, rice and petunia mutants with defects in SL biosynthesis or SL responses were first identified based on their increased shoot branching phenotypes and their dwarfism (Beveridge et al., 1996; Ishikawa et al., 2005; Napoli, 1996; Stirnberg et al., 2002). Less obvious phenotypes, such as reduced secondary growth, delay in leaf senescence or modified root architecture, were later identified (Brewer et al., 2013; Ueda and Kusaba, 2015; Yamada et al., 2014). SLs can also modulate tolerance to abiotic stresses (drought) (Ha et al., 2014). Direct or indirect roles for SLs in biotic stress-related responses have been suggested to act via crosstalk with other hormones (Al-Babili and Bouwmeester, 2015; Brewer et al., 2013; Stes et al., 2015). Thus, like other plant hormones, SLs can modulate multiple aspects of plant growth and development, either independently or via interactions with other hormonal and environmental pathways. The observed diversity of D53-like/SMXL proteins may contribute to the multiple processes controlled by SLs in plant development.

The origin and evolution of SL signaling

Studies have shown that species of the fresh water algae Nitella (Charales) are able to synthesize SLs (Delaux et al., 2012), suggesting that the SL pathway originated prior to the diversification of land plants (embryophytes). Since Charales do not establish symbiosis with AM fungi, it has been proposed that SLs first played a hormonal role during rhizoid elongation and were later recruited for symbiotic interactions (Delaux et al., 2012). In the moss Physcomitrella patens, SLs regulate protonema filament extension (Proust et al., 2011) as well as leafy shoot branching (Coudert et al., 2015). Furthermore, although SLs are detected in basal plants (Delaux et al., 2012), the KAI2/HTL clade appeared before the D14 clade, suggesting that D14 proteins might have been later selected as SL receptors during land plant evolution for novel developmental processes. Intriguingly, a high number of KAI2/HTL genes are present in the P. patens genome compared with angiosperms but also with Selaginella and Marchantia (Delaux et al., 2012). A similar KAI2/HTL gene expansion is found in parasitic plants (Conn et al., 2015; Tsuchiya et al., 2015). Interestingly, in these species, it was suggested that some of these KAI2/HTL homologs could be SL receptors (Conn et al., 2015). It should be noted that, despite SLs being detected in basal plants, the SL signaling pathway is poorly described in these plants. Recently, it was shown that KAI2/HTL homologs of Selaginella and Marchantia do not complement Arabidopsis d14 mutant phenotypes, nor some phenotypes of Arabidopsis kai2 (Waters et al., 2015). This leaves open the question of SL receptor identity in basal plants.

Perspectives

Despite significant progress, many key questions regarding SL biosynthesis, perception and signaling remain to be answered. The enzymatic activity of the SL receptor has been conserved during evolution, indicating that it plays an essential function, but this function is puzzling as the hydrolysis products (the ABC and D-OH parts) are known to be inactive. Is this enzymatic function of the SL receptor essential for SL reception and signaling? Does it play an important role in SL homeostasis? Furthermore, if SL perception truly involves an SL degradation process, are there other mechanisms of SL inactivation? The link between SL perception and downstream responses is also unclear. Although some downstream transcription factors have been identified, it is often noted that very few genes are transcriptionally regulated after SL application, at least over a short time frame, compared with other plant hormones (Smith and Li, 2014; Waldie et al., 2014), suggesting that non-transcriptional mechanisms might also be important in mediating the response to SL. Further investigation into both transcriptional and non-transcriptional responses and their importance will be key.

Understanding the molecular events acting downstream of D53/SMXL proteins will also be essential. In particular, it is unknown whether other post-translational modifications, such as phosphorylation and/or glycosylation, are required for the regulation of these downstream targets (Chen et al., 2014). The protein-protein interaction network in SL signaling also appears to be quite complex, and further understanding of these interactions might help to explain the observed crosstalk between the SL pathway and other plant signaling pathways. There is no doubt that the coming years will bring answers to the key questions in this exciting field.

Acknowledgements

We thank Alexandre de Saint Germain and Rajeev Kumar for comments on the manuscript. The IJPB benefits from the support of the Labex Saclay Plant Sciences-SPS [ANR-10-LABX-0040-SPS].

Footnotes

  • Competing interests

    The authors declare no competing or financial interests.

  • Funding

    We thank the Agence Nationale de la Recherche [contract ANR-12-BSV6-004-01] and the Stream COST Action FA1206 for financial support.

    Development at a Glance

    A high-resolution version of the poster is available for downloading in the online version of this article at http://dev.biologists.org/content/142/21/3615/F1.poster.jpg

  • © 2015. Published by The Company of Biologists Ltd

This is an Open Access article distributed under the terms of the Creative Commons Attribution License (http://creativecommons.org/licenses/by/3.0), which permits unrestricted use, distribution and reproduction in any medium provided that the original work is properly attributed.

References

Introduction

Strigolactones (SLs) are carotenoide-derived phytohormones that were originally identified as rhizosphere signal molecules, involved in parasitic and symbiotic interactions between plant roots and parasitic seeds/fungi (reviewed by Zhang et al., 2015). To date, more than 20 naturally occurring SL derivatives have been described (Al-Babili and Bouwmeester, 2015) fulfilling a plethora of roles in plant growth and development (reviewed by Obando et al., 2015). In 2008, SLs were identified as crucial regulators of plant branching (Gomez-Roldan et al., 2008; Umehara et al., 2008). In the following years it has been shown that SLs are also involved in regulating root development (Koltai and Kapulnik, 2014; Sun et al., 2016b), leaf senescence (Yamada and Umehara, 2015), and responses to nutrient stress (Marzec et al., 2013; Sun et al., 2016a), while a potential role in response to biotic stresses was recently proposed (Marzec and Muszynska, 2015).

Studies on mutant plants of Arabidopsis thaliana L., Oryza sativa L., Pisum sativum L., and Petunia hybrida L. enabled the identification of key proteins involved in SL biosynthesis and signaling. Biosynthesis of SL starts with the conversion of all-trans-β-carotene into carlactone (CL). This process takes place in plastids and involves a carotenoid isomerase and two carotenoid cleavage dioxygenases (Alder et al., 2012). Following its transport into the cytoplasm, MAX1-type monooxygenases transform CL into carlactonic acid, that is later converted into 5-deoxystrigol or orobanchol, two main precursors of other SLs (Seto et al., 2014). SLs consist of a tricyclic lactone (ABC ring) connected to a butenolide group (D ring). The C-D part is conserved among all SLs, while the A-B rings are subjected to modifications, including substitutions of the methyl, hydroxyl, and acetyloxyl groups (Figure 1). Based on the steric orientation of the α- (orobanchol-configured) or β-oriented (strigol-configured) C-ring SLs have been divided into two groups (Xie et al., 2013).

FIGURE 1.Structures of SLs. (A) Structure of strigol, the first identified representative of SLs, (B) structure of 5-deoxystrigol, the precursor of other β-oriented C-ring SLs (strigol-configured SLs), (C) structure of orobanchol, an example of SLs that carries an α-oriented C-ring (orobanchol-configured SLs), (D) structure of GR24, the synthetic analog of SLs.

In contrast to the biosynthesis pathway, knowledge about the SL signaling remained limited. Recent studies, however, brought great progress in uncovering the SL signaling mechanisms and components involved in SL perception, signal conversion and downstream responses in plants.

SL Perception

Analysis of SL-insensitive mutants enabled the identification of potential SL receptors in various plant species: D14 in rice (Arite et al., 2009), AtD14 in A. thaliana (Waters et al., 2012), DAD2 in petunia (Hamiaux et al., 2012), HvD14 in Hordeum vulgare L. (Marzec et al., 2016), and PtD14 in Populus trichocarpa Torr. & A. Gray (Zheng et al., 2016). All these receptors are members of the α/β-hydrolase family and are able to bind and hydrolyze SL molecules in vitro (Kagiyama et al., 2013; Nakamura et al., 2013). The enzymatic activity of the D14/DAD2 protein depends on the presence of the catalytic Ser/His/Asp triad (Hamiaux et al., 2012). In DAD2, substitution of the Ser96 by Ala resulted in a loss of catalytic activity and SL perception (Hamiaux et al., 2012). X-ray crystallography analysis of the D14/DAD2 protein revealed that the Ser within the catalytic triad is also involved in binding the D ring of SLs (Zhao et al., 2013). When the SL molecule is attached to the D14/DAD2, a nucleophilic attack separates the ABC part of the SL molecule from the D ring (Scaffidi et al., 2012). This reaction also results in a change of the D14/DAD2 conformation (Nakamura et al., 2013), which is crucial for the interaction of this protein with other components of the SL signaling complex (Zhao et al., 2015) (Figure 2). The binding pocket of D14/DAD2 is partially covered by a cap formed by four helicases (Kagiyama et al., 2013; Nakamura et al., 2013). Studies on the barley mutant hvd14.d revealed that the loss of function may be also due to a reduction of the aperture of entry to the binding pocket of the D14/DAD2 protein (Marzec et al., 2016). It has to be highlighted that the D14/DAD2 protein is a specific receptor for SLs, since karrikins and other regulators of plant growth and development that are structurally similar to SLs, are not recognized by this protein (Waters et al., 2012). The dynamics by which the D14/DAD2 receptor recognizes and hydrolyses different SL compounds, depends on the stereospecificity of SLs compounds (reviewed by Flematti et al., 2016) which thus play a crucial role in SLs perception and plant responses.

FIGURE 2.Scheme of the SL signaling pathway. (A) Expression of transcription factors (TFs) from the TCP family is repressed in the absence of SLs. (B) SL molecules are recognized by the SL receptor D14/DAD2. (C) The receptor hydrolyses SL molecules resulting in conformation changes of the D14/DAD2 protein. (D) In presence of SLs, the receptor with altered conformation is able to bind the F-box protein (MAX2/D3) from the SCF complex and the SL repressor (D53/SMXL6 to 8). (E) The repressor is degraded in the proteasome, also receptor is destabilized because of its changed conformation. (F) Degradation of repressor allows the expression of TFs from the TCP family. (G) List of identified components of the SLs signaling pathway in rice and A. thaliana.

Although AtD14 expression is found in all major plant organs, it still shows a high tissue specificity. For example in roots, expression of AtD14 was mainly in the vascular cylinder of the differentiation and elongation zones, whereas in leaves or cotyledons, a higher expression of AtD14 was observed in the phloem (Chevalier et al., 2014). Intriguingly, the pattern of AtD14 gene expression does not correspond to the AtD14 protein presence. For example, the AtD14 protein was found in nuclei of root meristem and rhizodermal cells, which were without relevant gene expression, indicating that either the mRNA or the D14/DAD2 protein is transported between the cells. Indeed, grafting studies confirmed that the D14/DAD2 protein is able to move between cells by short distance transport (Hamiaux et al., 2012; Chevalier et al., 2014).

Abundance of AtD14 mRNA did not change after treatment with auxin or the synthetic SL analog GR24, as well as during axillary bud development (Chevalier et al., 2014). It was therefore postulated that regulation of receptor abundance occurs at the protein level. Indeed, treatment of A. thaliana seedlings with GR24 resulted in a decreased AtD14 protein content (Chevalier et al., 2014). X-ray crystallography and hydrogen-deuterium exchange mass spectrometry (HDX) of the rice protein OsD14 and its conformational change after binding to GR24 molecules showed that binding to GR24 destabilizes the OsD14 (Zhao et al., 2015). This was the first indication of a phytohormone degrading its own receptor and affecting its own perception. It would be worth to investigate if this unexpected relation between signal molecule and receptor is indeed specific for SLs or whether it presents a more general mode of action among phytohormones.

SL Signal Conversion

Degradation of targeted proteins via the ubiquitin-proteasome pathway plays a crucial role in the signaling pathway of most phytohormones (Wang C. et al., 2015). The central element of this system is the SKP1-CULLIN-F-BOX complex (SCF). SL perception involves recognition and binding of target proteins by F-Box proteins which are subsequently bound by Skp1, before Cullin, the main structural component of the SCF complex, connects the complex to ubiquitin ligase (Larrieu and Vernoux, 2015). Since the F-box protein component renders specificity to the whole CSF complex, each hormone/signaling molecule may have its own exclusive F-box protein component. The protein recognized by the F-box protein is ubiquitinated thus marking it for proteasomal degradation.

In studies on the A. thaliana mutant max2 and the rice mutant d3 an F-box protein involved in SL signaling was identified that was also part of an SCF ubiquitin ligase protein complex (Stirnberg et al., 2002; Ishikawa et al., 2005). In A. thaliana MAX2 forms the SCF complex together with AtCullin1 and ARABIDOPSIS SERINE/THREONINE KINASE 1 (ASK1), whereas in rice the D3 protein interacts with OsCullin1 and ORYZA SATIVA SKP1-LIKE1/5/20 (OSK1/5/20) (Stirnberg et al., 2007; Zhao et al., 2014) (Figure 2). Similar to other components of the SLs signaling pathway, MAX2/D3 has a nuclear localization and the expression patterns of genes encoding this protein were similar to those observed for D14/DAD2 (Stirnberg et al., 2007; Zhao et al., 2014). The interaction between MAX2/D3 and D14/DAD2 was experimentally confirmed, and was shown to be promoted by the presence of SLs (Hamiaux et al., 2012; Zhao et al., 2014). Bimolecular fluorescence complementation analysis in rice protoplasts confirmed a GR24-mediated interaction between D3 and D14 within the nucleus (Zhao et al., 2014). The properties of this interaction which is mediated by SLs and depends on the SL concentration, is also affected by the SL stereoisomers involved (Zhao et al., 2015).

While certain components of the SL signaling pathway appear specific for SLs, the MAX2/D3 element is also involved in karrikin signal transduction. It is suggested that MAX2 may be part of different SCF complexes that are able to bind a range of substrates/repressors (Nelson et al., 2011). Observations in rice, where D3 interacts with at least three different OSKs, confirm the hypothesis that MAX2 can interact with multiple SCF complexes (Zhao et al., 2014). Moreover, it has been shown that MAX2 is also involved in the degradation of BRASSINAZOLE-RESISTANT1 (BES1), the transcriptional effector of the phytohormone class of brassinosteroids (Wang et al., 2013).

A phylogenetic analysis revealed similarity of MAX2/D3 to the auxin receptor TRANSPORT INHIBITOR RESPONSE1 (TIR1) (Dharmasiri et al., 2005) and the jasmonate receptor CORONATINE INSENSITIVE1 (COI1) (Sheard et al., 2010). Although there is no evidence that MAX2/D3 acts as a SL receptor, it cannot be excluded that this protein may recognize other signaling molecules, such as karrikins, since the A. thaliana max2 mutant showed a karrikin-resistant phenotype (Nelson et al., 2011).

All these data indicate that MAX2/D3 is probably involved in multiple signaling pathways and/or is a connector between SL perception and other phytohormones. This is a reason why to investigate the role of SLs in different aspects of plant growth and development it is better to use the SL-synthesis mutants or mutants in D14/DAD2 genes, that encoding receptor specific only for SLs. Whereas the results obtained for max2/d3 mutants might be related to their multiple role in plant signaling network. Now the identification of specific molecules recognized by MAX2/D3, as well as the identification of targets for the SCFMAX2/D3 complex is necessary to uncover the comprehensive role of this protein in the plant signaling network.

SL Signaling

The first SL repressor identified was D53 from rice (Zhou et al., 2013). Similar to other components of the SL signaling pathway D53 was discovered in a screening of SL-insensitive mutants displaying semi-dwarf phenotypes and higher number of tillers compared to their wild-type counterparts. Interestingly both, d53 mutants and wild-type plants overexpressing OsD53 showed increased branching, suggesting that the mutation in D53, i.e., a deletion of five amino acids, confers gain-of-function. The role of D53 in repressing the SL signal was confirmed by the lower number of tillers in d53 plants with reduced expression of D53 (Zhou et al., 2013). Recently, three orthologous of D53 identified in A. thaliana were also found to act as suppressors in SL signaling and named SUPRESSOR OF MAX2-LIKE6 to 8 (SMXL6 to 8) (Soundappan et al., 2015; Wang L. et al., 2015). First report indicated that all three genes function redundantly as shown by the fact that a reduced branching phenotype was only observed in the triple mutant smxl6/7/8 (Wang L. et al., 2015). However, recently it was shown that the presence of a stabilized form of SMXL7 under native promotor, resulted in a phenotype characteristic for SL mutants (Liang et al., 2016). Thus the question if all three repressors function redundantly remains still open.

The gene products of D53, SMXL6 to SMXL8 are localized in the nucleus. The presence of SL molecules was found to promote the interaction between these proteins and the receptor D14 (Zhou et al., 2013; Wang L. et al., 2015). At the same time SLs also induce fast proteasome-mediated degradation of D53 (Zhou et al., 2013), SMXL6 (Wang L. et al., 2015), and SMXL7 (Soundappan et al., 2015). Since degradation of D53 was not observed in d3, d14 and d53 mutants, it was concluded that the presence of the D3-D14-D53 complex is necessary for the degradation of SL repressors (Figure 2). Although interactions between D14/AtD14, D3/MAX2, and D53/SMXL6 to 8 have been confirmed, the interaction between SMXL6 and MAX2 does not require the presence of D14 and the interaction between SMXL6 and AtD14 does not require MAX2 (Wang L. et al., 2015).

The SL repressors found in rice and A. thaliana contain a highly conserved ethylene-responsive element binding factor-associated amphiphilic repression (EAR) motif of five amino acids (F/L-D-L-N-L). This motif has been postulated to interact with the transcriptional corepressors TOPLESS and TOPLESS-RELATED PROTEINS (TPR2) (Zhou et al., 2013; Ke et al., 2015; Soundappan et al., 2015) (Figure 2). Using a yeast-two hybrid and Co-Immunoprecipitation assays, Wang L. et al. (2015) were able to confirm the interaction between SMXL6 to 8 and TPR2 in vivo. In A. thaliana it was recently shown that SMXL7, D14, and MAX2 interact in the nucleus in an SL-dependent manner (Liang et al., 2016).

Presence of at least three SL-repressors in A. thaliana indicates a diverse regulation of the SLs signaling pathway and thus increasing the range of influences on different aspects of plant development. Studies on individual SMXLs and identification of genes regulated by SCF complexes containing different repressors, will confirm this hypothesis.

SL-Elicited Responses

The final confirmation that D53/SMXL6 to 8 act as SL repressors was provided by gene expression analysis. Until now only one class of transcription factors (TFs), the TEOSINTE BRANCHED1/CYCLOIDEA/PROLIFERATING CELL FACTO-R1 family (TCP), has been described as downstream component in SL signaling (Braun et al., 2012) (Figure 2). Representatives of TCP TFs have been found in rice (FC1, FINE CULM1) and A. thaliana (AtBRC1, BRANCHED1), and their expression has been observed in axillary buds. Both AtBRC1 and FC1 were upregulated after treatment with GR24, confirming their role in SL-mediated plant responses (Aguilar-Martínez et al., 2007; Minakuchi et al., 2010). Expression of AtBRC1 was down-regulated in SL-biosynthesis mutant max3 and SL-signaling mutant max2 but up-regulated in triple mutant smxl6/7/8 (Soundappan et al., 2015; Wang L. et al., 2015). Similar results were found for HB53, one of the known target genes of AtBRC1, which was elevated in smxl6/7/8 plants (Wang L. et al., 2015).

Knowledge on the interactions of SLs repressors and corepressors will allow to predict which TFs might be regulated by SLs, thus enabling a forecast to the plant response to SLs on the transcriptional level. Moreover, the comparative transcriptome analysis of individual smxl mutants might also reveal if all repressors function redundantly or not.

Common and Unique Features of SL Perception

Based on the genetic analysis of SL-insensitive mutants in rice and A. thaliana, three main players in SL signal transduction have already been identified: receptor D14/DAD2, repressor D53/SMXL6 to 8 and F-box protein MAX2/D3, which is a part of the SCF complex. The SL signaling pathway shares similarities with those of other phytohormones. D14/DAD2, the receptor of SLs resembles the gibberellin receptor GID1 (Griffiths et al., 2006; Arite et al., 2009; Hamiaux et al., 2012). Furthermore, proteasome-mediated degradation of the repressor by the SCF complex is a well-known mechanism of phytohormone-regulated gene expression (Wang L. et al., 2015).

Other aspects of the SLs signaling pathway seem very specific though. In contrast to the closely related gibberellin receptor GID1, the SL receptor D14/DAD2 is able to hydrolyse its receptor molecules (Hamiaux et al., 2012). Even more intriguing is that during the hydrolysis of SLs the conformation of D14 also changes which initiates the destabilization of this protein (Chevalier et al., 2014). Together with the degradation of D53/SMXL6 to 8 the perception of SLs constitutes a unique phenomenon among plant hormones involving the successive degradation of signal molecule, receptor, and downstream effector.

Author Contributions

The author confirms being the sole contributor of this work and approved it for publication.

Conflict of Interest Statement

The author declares that the research was conducted in the absence of any commercial or financial relationships that could be construed as a potential conflict of interest.

Acknowledgments

Author thanks Prof. Nicolaus von Wirén, Dr. Michael Melzer, and Dr. Twan Rutten for critical reading of the manuscript. Author is supported by scholarships founded by Foundation for Polish Science (START 067.2015) and Ministry of Science and Higher Education (424/STYP/10/2015 and DN/MOB/245/IV/2015).

References

Aguilar-Martínez, J. A., Poza-Carrión, C., and Cubas, P. (2007). Arabidopsis BRANCHED1 acts as an integrator of branching signals within axillary buds. Plant Cell 19, 458–472. doi: 10.1105/tpc.106.048934

CrossRef Full Text | Google Scholar

Al-Babili, S., and Bouwmeester, H. J. (2015). Strigolactones, a novel carotenoid-derived plant hormone. Annu. Rev. Plant Biol. 66, 161–186. doi: 10.1146/annurev-arplant-043014-114759

CrossRef Full Text | Google Scholar

Alder, A., Jamil, M., Marzorati, M., Bruno, M., Vermathen, M., Bigler, P., et al. (2012). The path from β-carotene to carlactone, a strigolactone-like plant hormone. Science 335, 1348–1351. doi: 10.1126/science.1218094

CrossRef Full Text | Google Scholar

Arite, T., Umehara, M., Ishikawa, S., Hanada, A., Maekawa, M., Yamaguchi, S., et al. (2009). d14, a strigolactone-insensitive mutant of rice, shows an accelerated outgrowth of tillers. Plant Cell Physiol. 50, 1416–1424. doi: 10.1093/pcp/pcp091

CrossRef Full Text | Google Scholar

Braun, N., de Saint Germain, A., Pillot, J. P., Boutet-Mercey, S., Dalmais, M., Antoniadi, I., et al. (2012). The pea TCP transcription factor PsBRC1 acts downstream of strigolactones to control shoot branching. Plant Physiol. 158, 225–238.

Google Scholar

Chevalier, F., Nieminen, K., Sánchez-Ferrero, J. C., Rodríguez, M. L., Chagoyen, M., Hardtke, C. S., et al. (2014). Strigolactone promotes degradation of DWARF14, an α/β hydrolase essential for strigolactone signaling in Arabidopsis. Plant Cell 26, 1134–1150. doi: 10.1105/tpc.114.122903

CrossRef Full Text | Google Scholar

Dharmasiri, N., Dharmasiri, S., and Estelle, M. (2005). The F-box protein TIR1 is an auxin receptor. Nature 435, 441–445. doi: 10.1038/nature03543

CrossRef Full Text | Google Scholar

Flematti, G. R., Scaffidi, A., Waters, M. T., and Smith, S. M. (2016). Stereospecificity in strigolactone biosynthesis and perception. Planta 243, 1361–1373. doi: 10.1007/s00425-016-2523-5

CrossRef Full Text | Google Scholar

Gomez-Roldan, V., Fermas, S., Brewer, P. B., Puech-Pagès, V., Dun, E. A., Pillot, J. P., et al. (2008). Strigolactone inhibition of shoot branching. Nature 455, 189–194. doi: 10.1038/nature07271

CrossRef Full Text | Google Scholar

Griffiths, J., Murase, K., Rieu, I., Zentella, R., Zhang, Z.-L., Powers, S. J., et al. (2006). Genetic characterization and functional analysis of the GID1 gibberellin receptors in Arabidopsis. Plant Cell 18, 3399–3414. doi: 10.1105/tpc.106.047415

CrossRef Full Text | Google Scholar

Hamiaux, C., Drummond, R. S., Janssen, B. J., Ledger, S. E., Cooney, J. M., Newcomb, R. D., et al. (2012). DAD2 is an α/β hydrolase likely to be involved in the perception of the plant branching hormone, strigolactone. Curr. Biol. 22, 2032–2036. doi: 10.1016/j.cub.2012.08.007

CrossRef Full Text | Google Scholar

Ishikawa, S., Maekawa, M., Arite, T., Onishi, K., Takamure, I., and Kyozuka, J. (2005). Suppression of tiller bud activity in tillering dwarf mutants of rice. Plant Cell Physiol. 46, 79–86. doi: 10.1093/pcp/pci022

CrossRef Full Text | Google Scholar

Kagiyama, M., Hirano, Y., Mori, T., Kim, S. Y., Kyozuka, J., Seto, Y., et al. (2013). Structures of D14 and D14L in the strigolactone and karrikin signaling pathways. Genes Cells 18, 147–160. doi: 10.1111/gtc.12025

CrossRef Full Text | Google Scholar

Ke, J., Ma, H., Gu, X., Thelen, A., Brunzelle, J. S., Li, J., et al. (2015). Structural basis for recognition of diverse transcriptional repressors by the TOPLESS family of corepressors. Sci. Adv. 1, e1500107. doi: 10.1126/sciadv.1500107

CrossRef Full Text | Google Scholar

Koltai, H., and Kapulnik, Y. (2014). “Strigolactones involvement in root development and communications,” in Root Engineering, eds A. Morte and A. Varma (Berlin: Springer), 203–219.

Google Scholar

Larrieu, A., and Vernoux, T. (2015). Comparison of plant hormone signalling systems. Essays Biochem. 58, 165–181. doi: 10.1042/bse0580165

CrossRef Full Text | Google Scholar

Liang, Y., Ward, S., Li, P., Bennett, T., and Leyser, O. (2016). SMAX1-LIKE7 signals from the nucleus to regulate shoot development in Arabidopsis via partially EAR motif-independent mechanisms. Plant Cell 28, 1581–1601. doi: 10.1105/tpc.16.00286

CrossRef Full Text | Google Scholar

Marzec, M., Gruszka, D., Tylec, P., and Szarejko, I. (2016). Identification and functional analysis of the HvD14 gene involved in strigolactone signalling in Hordeum vulgare L. Physiol. Plant doi: 10.1111/ppl.12460 [Epub ahead of print].

CrossRef Full Text | Google Scholar

Marzec, M., and Muszynska, A. (2015). In silico analysis of the genes encoding proteins that are involved in the biosynthesis of the RMS/MAX/D pathway revealed new roles of strigolactones in plants. Int. J. Mol. Sci. 16, 6757–6782. doi: 10.3390/ijms16046757

CrossRef Full Text | Google Scholar

Marzec, M., Muszynska, A., and Gruszka, D. (2013). The role of strigolactones in nutrient-stress responses in plants. Int. J. Mol. Sci. 14, 9286–9304. doi: 10.3390/ijms14059286

CrossRef Full Text | Google Scholar

Minakuchi, K., Kameoka, H., Yasuno, N., Umehara, M., Luo, L., Kobayashi, K., et al. (2010). FINE CULM1 (FC1) works downstream of strigolactones to inhibit the outgrowth of axillary buds in rice. Plant Cell Physiol. 51, 1127–1135. doi: 10.1093/pcp/pcq083

CrossRef Full Text | Google Scholar

Nakamura, H., Xue, Y. L., Miyakawa, T., Hou, F., Qin, H. M., Fukui, K., et al. (2013). Molecular mechanism of strigolactone perception by DWARF14. Nat. Commun. 4, 2613. doi: 10.1038/ncomms3613

CrossRef Full Text | Google Scholar

Nelson, D. C., Scaffidi, A., Dun, E. A., Waters, M. T., Flematti, G. R., Dixon, K. W., et al. (2011). F-box protein MAX2 has dual roles in karrikin and strigolactone signaling in Arabidopsis thaliana. Proc. Natl. Acad. Sci. U.S.A. 108, 8897–8902. doi: 10.1073/pnas.1100987108

CrossRef Full Text | Google Scholar

Obando, M. L., Ligerot, Y., Boyer, F. D., and Rameau, C. (2015). Strigolactone biosynthesis and signaling in plant development. Development 142, 3615–3619. doi: 10.1242/dev.120006

CrossRef Full Text | Google Scholar

Scaffidi, A., Waters, M. T., Bond, C. S., Dixon, K. W., Smith, S. M., Ghisalberti, E. L., et al. (2012). Exploring the molecular mechanism of karrikins and strigolactones. Bioorg. Med. Chem. Lett. 22, 3743–3746. doi: 10.1016/j.bmcl.2012.04.016

CrossRef Full Text | Google Scholar

Seto, Y., Sado, A., Asami, K., Hanada, A., Umehara, M., Akiyama, K., et al. (2014). Carlactone is an endogenous biosynthetic precursor for strigolactones. Proc. Natl. Acad. Sci. U.S.A. 111, 1640–1645. doi: 10.1073/pnas.1314805111

CrossRef Full Text | Google Scholar

Sheard, L. B., Tan, X., Mao, H., Withers, J., Ben-Nissan, G., Hinds, T. R., et al. (2010). Jasmonate perception by inositol-phosphate-potentiated COI1-JAZ co-receptor. Nature 468, 400–405. doi: 10.1038/nature09430

CrossRef Full Text | Google Scholar

Soundappan, I., Bennett, T., Morffy, N., Liang, Y., Stanga, J. P., Abbas, A., et al. (2015). SMAX1-LIKE/D53 family members enable distinct MAX2-dependent responses to strigolactones and karrikins in Arabidopsis. Plant Cell 27, 3143–3159. doi: 10.1105/tpc.15.00562

CrossRef Full Text | Google Scholar

Stirnberg, P., Furner, I. J., and Leyser, O. (2007). MAX2 participates in an SCF complex which acts locally at the node to suppress shoot branching. Plant J. 50, 80–94. doi: 10.1111/j.1365-313X.2007.03032.x

CrossRef Full Text | Google Scholar

Stirnberg, P., van De Sande, K., and Leyser, H. O. (2002). MAX1 and MAX2 control shoot lateral branching in Arabidopsis. Development 129, 1131–1141.

Google Scholar

Sun, H., Bi, Y., Tao, J., Huang, S., Hou, M., Xue, R., et al. (2016a). Strigolactones are required for nitric oxide to induce root elongation in response to nitrogen and phosphate deficiencies in rice. Plant Cell Environ. 39, 1473–1483. doi: 10.1111/pce.12709

CrossRef Full Text

Sun, H., Tao, J., Gu, P., Xu, G., and Zhang, Y. (2016b). The role of strigolactones in root development. Plant Signal. Behav. 11, e1110662. doi: 10.1080/15592324.2015.1110662

CrossRef Full Text | Google Scholar

Umehara, M., Hanada, A., Yoshida, S., Akiyama, K., Arite, T., Takeda-Kamiya, N., et al. (2008). Inhibition of shoot branching by new terpenoid plant hormones. Nature 455, 195–200. doi: 10.1038/nature07272

CrossRef Full Text | Google Scholar

Wang, C., Liu, Y., Li, S. S., and Han, G. Z. (2015). Insights into the origin and evolution of the plant hormone signaling machinery. Plant Physiol. 167, 872–886. doi: 10.1104/pp.114.247403

CrossRef Full Text | Google Scholar

Wang, L., Wang, B., Jiang, L., Liu, X., Li, X., Lu, Z., et al. (2015). Strigolactone signaling in Arabidopsis regulates shoot development by targeting D53-like SMXL repressor proteins for ubiquitination and degradation. Plant Cell 27, 3128–3142. doi: 10.1105/tpc.15.00605

CrossRef Full Text | Google Scholar

Wang, Y., Sun, S., Zhu, W., Jia, K., Yang, H., and Wang, X. (2013). Strigolactone/MAX2-induced degradation of brassinosteroid transcriptional effector BES1 regulates shoot branching. Dev. Cell 27, 681–688. doi: 10.1016/j.devcel.2013.11.010

CrossRef Full Text | Google Scholar

Waters, M. T., Nelson, D. C., Scaffidi, A., Flematti, G. R., Sun, Y. K., Dixon, K. W., et al. (2012). Specialisation within the DWARF14 protein family confers distinct responses to karrikins and strigolactones in Arabidopsis. Development 139, 1285–1295. doi: 10.1242/dev.074567

CrossRef Full Text | Google Scholar

Xie, X., Yoneyama, K., Kisugi, T., Uchida, K., Ito, S., Akiyama, K., et al. (2013). Confirming stereochemical structures of strigolactones produced by rice and tobacco. Mol. Plant 6, 153–163. doi: 10.1093/mp/sss139

CrossRef Full Text | Google Scholar

Yamada, Y., and Umehara, M. (2015). Possible roles of strigolactones during leaf senescence. Plants 4, 664–677. doi: 10.3390/plants4030664

CrossRef Full Text | Google Scholar

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