, Volume 243, Issue 6, pp 1339–1350 | Cite as

Strigolactone versus gibberellin signaling: reemerging concepts?

  • Eva-Sophie Wallner
  • Vadir López-Salmerón
  • Thomas GrebEmail author
Open Access
Part of the following topical collections:
  1. Strigolactones


Main conclusion

In this review, we compare knowledge about the recently discovered strigolactone signaling pathway and the well established gibberellin signaling pathway to identify gaps of knowledge and putative research directions in strigolactone biology.

Communication between and inside cells is integral for the vitality of living organisms. Hormonal signaling cascades form a large part of this communication and an understanding of both their complexity and interactive nature is only beginning to emerge. In plants, the strigolactone (SL) signaling pathway is the most recent addition to the classically acting group of hormones and, although fundamental insights have been made, knowledge about the nature and impact of SL signaling is still cursory. This narrow understanding is in spite of the fact that SLs influence a specific spectrum of processes, which includes shoot branching and root system architecture in response, partly, to environmental stimuli. This makes these hormones ideal tools for understanding the coordination of plant growth processes, mechanisms of long-distance communication and developmental plasticity. Here, we summarize current knowledge about SL signaling and employ the well-characterized gibberellin (GA) signaling pathway as a scaffold to highlight emerging features as well as gaps in our knowledge in this context. GA signaling is particularly suitable for this comparison because both signaling cascades share key features of hormone perception and of immediate downstream events. Therefore, our comparative view demonstrates the possible level of complexity and regulatory interfaces of SL signaling.


D53/SMXL Hormonal signaling Long-distance communication SCF complex 









SLs have a long research history in the context of interactions between plants and other organisms. They were identified in 1966 as plant-derived molecules used by parasitic plants to interact with their hosts (Cook et al. 1966). Further emphasizing their importance for biotic interactions, the role of SLs in the establishment of symbioses between plants and arbuscular mycorrhizal fungi was revealed in 2005 (Akiyama et al. 2005). Only in 2008 were SLs recognized as endogenous phytohormones when their role as decisive hormones regulating plant architecture was uncovered (Gomez-Roldan et al. 2008; Umehara et al. 2008). Since then, research on SL signaling mechanisms has revealed surprising parallels to other hormone signaling cascades, with the most similar being mechanisms of GA perception. Due to the instructive nature of comparative approaches, we relate in this review GA and SL signaling in order to accentuate emerging similarities and differences between the two pathways. Due to the striking parallels between both signaling cascades, we hope that this approach will be helpful for understanding the biological role of SL signaling during plant growth. For example, the presence of different bioactive GAs or the parallel effects of GA on transcription and subcellular localization of proteins demonstrates the complexity of molecular events that should be considered for a comprehensive understanding of a hormonal signaling cascade.

It is important to note, however, that there is no reason to think that SL signaling is more entangled with GA signaling than with other hormonal signaling pathways. Indeed, the interaction between auxin and SL signaling has a long history of research (Waldie et al. 2014; Brewer et al. 2009, 2015; Domagalska and Leyser 2011). Furthermore, the concept that nuclear hormone receptors, inducing the degradation of signaling repressors, extensively discussed in this review, is not restricted to GA and SL signaling but also found in jasmonic acid and auxin signaling cascades (Larrieu and Vernoux 2015). However, for the sake of conciseness we focus on the GA-SL comparison in order to guide the potential routes of SL research and demonstrate gaps in current knowledge. For the same reason, we do not focus on mechanisms of GA or SL biosynthesis, although this is an essential level of regulation, as this has been recently presented in excellent and comprehensive overviews (Seto and Yamaguchi 2014; Hedden and Thomas 2012).

Similar but different—families of related molecules

More than 100 different GAs have been isolated from vascular plants (MacMillan 2001) from which gibberellin A1 (GA1), GA3, GA4, GA5, GA6 and GA7 are biologically active. These GAs show different affinities to their receptors (Ueguchi-Tanaka et al. 2005, 2007; Nakajima et al. 2006) and their occurrence and abundance varies between different plant species (MacMillan 2001). For example, whereas GA1 is the most widespread gibberellin among species, GA4 is the most abundant and relevant bioactive GA in Arabidopsis (Eriksson et al. 2006; Talon et al. 1990). The structural requirements for a bioactive GA are clearly defined. These diterpenoid acids must possess a carboxyl group at position C6, a hydroxyl group at position C3 in β-orientation and a γ-lactone ring. Furthermore, they must not be hydroxylated at position C2, since hydroxylation at this position is critical for inactivation of GA in planta (Ueguchi-Tanaka and Matsuoka 2010) (Fig. 1). The stability of different GAs is also important to consider. GA3, for instance, shows a lower affinity than GA4 to its receptor GIBBERELLIN-INSENSITIVE DWARF1 (GID1) but a higher bioactivity. This is presumably due to increased GA3 stability caused by a double bond at the C2 position (Ueguchi-Tanaka et al. 2005).
Fig. 1

Similarities between SL and GA perception. a Molecular structures of SL and GA are exemplified by (+)-5-Deoxystrigol and GA3, respectively. The ABC scaffold of SL is connected to ring D by an enol ether bridge (indicated in orange). b A schematic comparison between SL- and GA-signaling is shown. Unlike GID1, the α/β-hydrolase D14 preserved its catalytic activity. Bound SL is hydrolyzed through a nucleophilic attack by Ser147 (visualized in orange) at the enol ether bridge. Marvin was used for drawing, displaying and characterizing chemical structures, substructures and reactions, Marvin Beans (, 2015, ChemAxon ( Abbreviations, see main text

Although identification of SLs is technically very challenging, around 20 naturally occurring SLs have been described so far (Zwanenburg and Pospisil 2013; Ueno et al. 2014). They all share an ABC scaffold consisting of three carbon rings attached to a butenolide (ring D) by an enol ether bridge (Fig. 1) (Zwanenburg et al. 2015; Xie and Yoneyama 2010). The enol ether bridge determines the bioactivity of SLs, since hydrolytic cleavage between ring C and D is crucial for SL perception and specificity (Zwanenburg et al. 2013, 2015; Mangnus and Zwanenburg 1992). The importance of the CD rings becomes obvious by the finding that an additional methyl group on ring D can significantly decrease the molecule’s ability to induce parasitic seed germination (Zwanenburg et al. 2013). Depending on the stereochemistry of the BC junction, SLs fall into the strigol and orobanchol classes, which show an opposing C-ring orientation determining functional specificity (Zwanenburg et al. 2015; Zhang et al. 2014; Scaffidi et al. 2014). 5-Deoxystrigol (5DS) and 4-deoxyorobanchol (4DS) are most likely the parent molecules that are converted into both classes, respectively, with overlapping but not identical biological activities (Zhang et al. 2014; Zwanenburg et al. 2015; Scaffidi et al. 2014). For instance, members of the strigol class most effectively stimulate germination of the parasitic weed Striga hermonthica, whereas orobanchol derivatives show the highest activity in stimulating mycorrhizal hyphal branching (Nomura et al. 2013; Akiyama et al. 2010). Beside these canonical SLs, a major role of non-canonical SLs, like methyl carlactonate, has been discussed especially for Arabidopsis (Abe et al. 2014; Zhang et al. 2014).

It is important to note that, although the bioactivity of individual SLs and in vitro receptor binding was shown in some cases (see below), the identification of the active forms in planta is a challenging enterprise. This is, in part, because plants may quickly convert applied compounds. A deeper understanding of the SL biosynthetic pathway and analysis of respective mutants will be essential to clarify which features are crucial for bio-availability of naturally occurring SLs (Seto and Yamaguchi 2014). For example, in addition to 2β-hydroxylation, bioactive GAs are also inactivated by methylation (Varbanova et al. 2007) and epoxidation of the C-16,17 double bond (Zhu et al. 2006). GA-methyl transferase activity mediated by GIBBERELLIN METHYLTRANSFERASE1 (GAMT1) and GAMT2 appears to be restricted to developing seeds (Varbanova et al. 2007; Nir et al. 2014) whereas 16,17-epoxidation has only been demonstrated in rice (Zhu et al. 2006). In the case of SLs it has not been determined if there are essential regulatory modulations of bioactive SL molecules.

Due to the high variability and specificity within the SL family, artificially produced SL analogs of simplified structure have to be used cautiously (Conn et al. 2015; Zwanenburg et al. 2015). Plants do not produce these analogs, which may, therefore, act very differently from endogenous SLs. For instance, the synthetic and broadly used SL analog GR24 consists of a racemic mixture of natural strigol-like GR245DS as well as its unnatural enantiomer GR24ent−5DS (Scaffidi et al. 2014; Conn et al. 2015). The natural GR245DS is most active in repressing SL-dependent shoot branching, whereas GR24ent−5DS preferentially activates the karrikin (KAR)-dependent pathway inducing germination after wildfires (Conn et al. 2015; Umehara et al. 2015; Waters et al. 2014) and important for recruiting arbuscular mycorrhizal fungi in rice (Gutjahr et al. 2015). Therefore, the effects observed after GR24 application are not necessarily natural SL responses.

It has not been reported that different bioactive GAs trigger different responses (Nakajima et al. 2006). All GID1 family members display a similar profile of binding affinities (Nakajima et al. 2006). This is interesting, as triggering specific subsets of downstream responses by different GAs could provide an advantage by providing regulatory flexibility. However, GAs are not only produced by plants but also by fungal pathogens to manipulate plant growth (Bömke and Tudzynski 2009). Prevention of sophisticated growth manipulation by pathogens may be a reason for this lack of signaling complexity among GA molecules. Although KAR receptors may sense fungus-derived signals (see below) (Gutjahr et al. 2015), there is no indication that non-plant pathogens produce SLs. The more complex set of SL-related molecules may be important for the recruitment of host- and/or growth stage-specific sets of symbiotic fungi (Gutjahr 2014) on the one side and the avoidance of parasitic plants (Cardoso et al. 2011) on the other side. Therefore, a spectrum of different SLs with slightly different activity is likely to be under positive selection (Akiyama et al. 2010; Nomura et al. 2013). The presence of canonical SLs in rice, which hosts mycorrhizal fungi, and their apparent absence in the non-host plant Arabidopsis (Abe et al. 2014) may be an example for species-specific adaptation.

The importance of hormone distribution

GAs move over long distances (Ragni et al. 2011; Proebsting et al. 1992) and recently it was suggested that GA12, the precursor of bioactive GAs, is the main form traveling along the vasculature (Regnault et al. 2015). Importantly, the finding that fluorescently labelled and bioactive GAs accumulate particularly in the root endodermis suggests that differential accumulation of GAs in plants occurs (Shani et al. 2013). The endodermis is also the most potent tissue for influencing GA-dependent root elongation (Ubeda-Tomas et al. 2008) and a site for GA production (Zhang et al. 2011). Overall, the fundamental role of spatial regulation of hormone levels and signaling is an emerging picture in many contexts (Savaldi-Goldstein et al. 2007; Iyer-Pascuzzi et al. 2011) and is especially established for auxins (Adamowski and Friml 2015).

The spatial distribution of SLs has not been revealed with high resolution; but novel fluorescent and bioactive SL analogs may provide an angle for filling this gap of knowledge (Prandi et al. 2013; Rasmussen et al. 2013b; Artuso et al. 2015; Fridlender et al. 2015). The expression of SL biosynthesis genes is usually highest in roots and partially associated with vascular tissues (Booker et al. 2005; Kohlen et al. 2012). Indeed, SL-like bioactivity has been found in the Arabidopsis xylem sap (Kohlen et al. 2011). Although an important role of canonical SLs in Arabidopsis was questioned in later studies (Abe et al. 2014), orobanchol was identified directly in the tomato and Arabidopsis xylem sap, pointing out a possibility for long-distance movement (Kohlen et al. 2011, 2012). In any case, movement of SLs—or their precursors—is able to completely suppress effects of SL-deficiency in grafting experiments with a preferred directionality for traveling from roots to shoots (Foo and Davies 2011; Turnbull et al. 2002; Booker et al. 2005). The low pH usually found in the xylem sap (Jia and Davies 2007) would support SL stability (Zwanenburg et al. 2015). Candidates for moving long distances are carlactonoic acid and orobanchol, the suggested products of MORE AXILLARY GROWTH1 (MAX1)-like enzymes, which catalyze the last step in the SL biosynthetic chain (Abe et al. 2014; Zhang et al. 2014; Booker et al. 2005). Consequently, the diverse regulatory roles of SLs, such as inhibiting shoot branching, promoting cambium activity and regulating root growth, partly in response to environmental cues (Umehara et al. 2010), would provide a means for coordinating plant growth processes in a systemic manner (Agustí et al. 2011; Gomez-Roldan et al. 2008; Rasmussen et al. 2013a; Umehara et al. 2010). However, the relevance of hormone movement under natural conditions is difficult to demonstrate without a possibility to manipulate this movement in a very specific manner. The lack of knowledge on how GA travels through the plant has hampered research in this direction so far. The discovery that the ABC transporter PLEIOTROPIC DRUG RESISTANCE1 (PDR1) from petunia (Petunia axillaris) is involved in SL secretion into the rhizosphere (Kretzschmar et al. 2012) and localizes polarly in plasma membranes (Sasse et al. 2015) may provide a novel avenue in this context. Thus, in addition to a passive long-distance movement, mechanisms for establishing local SL maxima may exist, which are relevant for local and cell type-specific responses.

The conversion of enzymes into receptors

The most striking analogy between GA and SL signaling is the mechanism of perception. The nuclear-localized and soluble protein GID1 is a catalytically inactive α/β-hydrolase identified in rice, which binds bioactive GAs (Ueguchi-Tanaka et al. 2005; Shimada et al. 2008). In comparison to rice, which possesses only one GID1 gene, there are three redundant GID1 genes (GID1a,b and −c) in Arabidopsis (Nakajima et al. 2006; Griffiths et al. 2006). Single mutants show only mild phenotypic alterations, but the gid1a/b/c triple mutant displays an extremely dwarfed growth habit and complete GA insensitivity (Griffiths et al. 2006; Ueguchi-Tanaka et al. 2005). This indicates that these proteins are the only GA receptors. The crystal structure of the GID1 receptor has helped to understand its function and the structural requirements that define a bioactive GA (Shimada et al. 2008). GA binding triggers a conformational change in the GID1 protein. This change promotes direct interaction of the GA-GID1 complex with DELLA proteins acting as transcriptional regulators (Harberd et al. 2009; Sun 2011). Formation of the GA-GID-DELLA ternary complex, in turn, recruits the SCFSLY1 (SKP1, CULLIN, F-box and RBX1 RING-domain) ubiquitin ligase (E3) complex via the F-box protein SLEEPY1 (SLY1), which provides substrate specificity to the complex (Dill et al. 2004) (Fig. 1).

As described below, physical contact of DELLA proteins with the SCFSLY1 complex results in their ubiquitination and degradation by the 26S proteasome (Harberd et al. 2009; Dill et al. 2004). Removal of the nuclear DELLA proteins results in massive changes in gene expression and, among other things, culminates in cell elongation (Harberd et al. 2009). In this respect, it is remarkable that sly1 mutants (or gid2 mutants in rice) show much milder phenotypic alterations than gid1a/b/c mutants do, although they accumulate comparable or even higher levels of DELLAs. Intriguingly, overexpression of the GID1 receptor suppresses these alterations (Ariizumi et al. 2008; Ueguchi-Tanaka et al. 2008). Thus, GID1 proteins may also play a GA-independent role in modulating DELLA activity, by sequestering these repressors into an inactive complex (Ariizumi et al. 2008; Ueguchi-Tanaka et al. 2008; Hauvermale et al. 2014).

In analogy to GA perception, substantial evidence has been provided that SLs bind to the α/β hydrolase DWARF14/DECREASED IN APICAL DOMINANCE2 (D14/DAD2) (Kagiyama et al. 2013; Zhou et al. 2013; Hamiaux et al. 2012). The binding pocket of D14/DAD2 contains the catalytic tirade Ser147, Asp268 and His297, which hydrolyzes the enol ether bridge between the C and D ring through a nucleophilic attack by Ser147 (Fig. 1) (Kagiyama et al. 2013; Zhao et al. 2015). Any similar activity has been lost in GID1 due to an amino acid substitution that replaced His by Val (Ueguchi-Tanaka et al. 2005). Because reaction products of D14/DAD2 do not display any biological activity, the decisive step in signal transduction is the conformational change of the D14/DAD2 protein and not the generation of signaling molecules (Hamiaux et al. 2012). D14/DAD2 is homologous to the KAR receptor KARRIKIN INSENSITIVE2 (KAI2). However, structure determination and binding analyses revealed that only D14/DAD2 binds SLs (Guo et al. 2013; Conn et al. 2015; Nakamura et al. 2013; Hamiaux et al. 2012; Toh et al. 2015; Zhao et al. 2015). In fact, it seems as if diversification of SL receptor-like proteins was crucial for the establishment of these distinct signaling cascades (Conn et al. 2015; Waters et al. 2012), a situation not found in the case of GID1. In addition to mediating KAR-dependent seed germination in some species, still unknown endogenous KAI2-binding molecules must exist because kai2 mutants display also developmental defects (Nelson et al. 2011; Waters et al. 2012). Interestingly, the KAI2 ortholog D14L in rice is essential for the recognition of arbuscular mycorrhizal fungi and the initiation of symbiotic interactions (Gutjahr et al. 2015). Thus, SL/KAR-related molecules do not only act as attractants during biotic interactions but their endogenous perception machinery is also important for recruiting symbiotic organisms. This argues for an intensive SL/KAR-dependent cross talk bridging species boundaries. The existence of a third D14/DAD2-like protein in Arabidopsis designated as D14-LIKE2 (DLK2), which does not contribute to SL or KAR responsiveness (Waters et al. 2012), suggests an even more complex situation on this level.

Similar to GID1, D14/DAD2 changes conformation upon SL binding which facilitates the interaction with the F-box protein and SCF complex component DWARF3 (D3). D3 is the rice ortholog to MORE AXILLARY GROWTH2 (MAX2) from Arabidopsis which is mainly expressed in vascular tissues (Chevalier et al. 2014; de Saint et al. 2013a, b; Zhou et al. 2013; Jiang et al. 2013; Stirnberg et al. 2007). Binding of D3/MAX2 to D14/DAD2 occurs close to its lid domain (Zhao et al. 2015) (Fig. 1). In comparison to SLY1, which can be partly replaced by the F-box protein SNEEZY (SNE) (Ariizumi et al. 2011), D3/MAX2 is the only F-box protein known to act in SL signaling. In fact, D3/MAX2 plays a key role in both the D14/DAD2 and KAI2-dependent signaling pathways (Waters et al. 2012). Interestingly, an exclusive role of D3/MAX2 in SL/KAR-signaling is questioned by the observation that max2 mutants respond to higher GR24 concentrations (Ruyter-Spira et al. 2011; Agustí et al. 2011) for which the basis still has to be determined. As explained in more detail below, the SCFD3/MAX2 E3 ubiquitin ligase complex executes SL-dependent ubiquitination of target proteins, such as DWARF53 (D53) in rice (Jiang et al. 2013). Just as the ubiquitination machinery of GA signaling and its DELLA targets, D3/MAX2, D14 and D53 are nuclear localized (Jiang et al. 2013; Stirnberg et al. 2007; Nakamura et al. 2013), thereby providing a potential link to a direct regulation of gene transcription.

Of note, GA and SL signaling pathways have been suggested to directly interact with each other. Hydrolyzation of SL/GR24 enables D14 to bind not only to D53-like proteins but also SLENDER1 (SLR1), the only DELLA protein found in rice (Nakamura et al. 2013). Thereby, SLs may contribute to GA signaling and suppress bud outgrowth in rice (Nakamura et al. 2013). However, D14-SLR1 binding was only shown indirectly using heterologous expression systems, and there is no physiological or genetic evidence that both pathways intertwine functionally. Instead, there are indications favoring an independent action. SL signaling promotes internode elongation in peas by increasing cell number, not by stimulating cell elongation as primarily done by GA (de Saint et al. 2013b). Furthermore, GA, but not GR24, application destabilizes DELLA proteins, GA responsiveness is not affected in SL signaling mutants and their dwarfism is not correlated with reduced GA levels (de Saint et al. 2013b). Further supporting an independent action, SL signaling acts antagonistically rather than in concert with GA signaling in the regulation of shoot branching in the woody plant Jatropha curcas (Ni et al. 2015).

Direct targets of signaling—the reemerging motif of repressing repressors

As mentioned, binding of GA or SLs to their respective receptor complexes leads to the 26S proteasome-dependent degradation of two distinct groups of signaling repressors: DELLA proteins in the case of GA and D53-like proteins in the case of SLs (Jiang et al. 2013; Zhou et al. 2013). DELLA proteins belong to the larger family of GRAS transcriptional regulators, which seem to have diversified to allow the integration of GA signaling into transcriptional regulation. DELLA proteins are named after their N-terminally conserved amino acid sequence (D–E–L–L–A) essential for binding to GID1 (Schwechheimer and Willige 2009; Wang and Deng 2011). In Arabidopsis, the GRAS proteins GA-INSENSITIVE (GAI), REPRESSOR OF GA1-3 (RGA), RGA-LIKE1 (RGL1), RGL2 and RGL3 carry such a domain (Dill et al. 2004). Although partially redundant, the five DELLA proteins display a certain functional specialization, such as the regulation of germination, stem elongation, leaf expansion, apical dominance or floral development (Dill et al. 2004; Wang and Deng 2011). While this specialization appears to result rather from their distinct expression patterns than from differences in protein properties (Gallego-Bartolome et al. 2010), there is an indication that there are differences in GA-induced degradation kinetics among the DELLA proteins (Wang et al. 2009) although more accurate studies are required to confirm these differences. Interestingly, in contrast to SLY1 which targets all DELLAs equally, SNE preferentially targets RGA and GAI, thus providing the possibility for a differential regulation of DELLA protein abundance on the level of the GA perception machinery (Ariizumi et al. 2011).

The reasonable assumption that SL signaling depends on the proteolysis of a set of repressor proteins was confirmed by the seminal identification of the D53 protein in rice which is nuclear localized and shows weak similarities to Class 1 Hsp100/ClpB proteins (Jiang et al. 2013; Zhou et al. 2013). Reminiscent of the GA-effect on DELLA proteins, D53 interacts with both D3 and D14 in an SL-dependent manner and is subsequently ubiquitinated and degraded (Jiang et al. 2013; Zhou et al. 2013). The d53 rice mutant carries a dominant-negative allele producing a protein with a deletion of five amino acids (GKTGI) and an amino acid substitution that changes a positively charged Arg into a Thr (Fig. 2). This alteration results in GR24-insensitivity and a dwarfed and bushy phenotype indicative of reduced SL signaling (Jiang et al. 2013; Zhou et al. 2013). Although both D53 and the mutated d53 protein are able to interact with D14, only D53 undergoes SL-dependent proteolysis (Jiang et al. 2013). This indicates that, unlike the DELLA motif, the RGKTGI sequence is crucial for the D14–D3-complex dependent ubiquitination but not for the interaction with the SL receptor complex. In fact, the part of D53-like proteins that interacts with the D14–D3 complex is still to be determined.
Fig. 2

Comparison of D53/SMXL family members. a A maximum likelihood phylogenic tree based on an amino acid sequence alignment of the Arabidopsis SMXL proteins. The scale bar indicates a branch length with 0.5 amino acid substitutions per site. The three putative sub-clades are emphasized by blue brackets. CLC Main Workbench 7.6.1 (CLC Bio Qiagen, Denmark). b Shown is the motif important for D3-dependent ubiquitination of D53 from rice identified previously (Jiang et al. 2013; Zhou et al. 2013). Aligned are the eight SMXL family members from Arabidopsis, the SMXL rice homolog D53 (OsD53) and the mutated d53 protein in which this motif is lost (indicated by a red bracket). Note that the RGKTGI motif is not present in members of sub-clade 2. CLC Main Workbench 7.6.1 (CLC Bio Qiagen, Denmark)

Consistent with the idea that the SL signaling mechanism is conserved across species boundaries, the D53 homologue SUPPRESSOR OF MAX2 1 (SMAX1) was identified in Arabidopsis in an elegant forward genetic screen for suppressors of effects of impaired SL/KAR signaling (Stanga et al. 2013). SMAX1 defined the small gene family of SMAX1-LIKE (SMXL) proteins consisting of eight members in Arabidopsis (Fig. 2). Similar to the DELLA proteins, differences in specificity and function have been proposed for SMXL family members (Stanga et al. 2013). The smax1 max2 mutant suppresses hypocotyl and germination defects found in max2 mutants, but not the typical increase in shoot branching, which is primarily associated with SL-deficiency (Stanga et al. 2013). Because SMAX1 and SMXL2, the two members of the D53/SMXL sub-clade 1, are sufficient for regulating all KAR-dependent responses, a functional separation of the D53/SMXL family into KAR and SL-signaling factors is likely (Stanga et al. 2013, 2016; Waters et al. 2014). Consistent with this idea, triple mutants lacking the activity of the clade 3-family members, SMXL6, SMXL7 and SMXL8, fully suppress all SL-related growth alterations caused by MAX2-deficiency (Wang et al. 2015; Soundappan et al. 2015). As with D53 in rice, the nuclear-localized SMXL6, SMXL7 and SMXL8 proteins are ubiquitinated and degraded upon the addition of GR24. Likewise, they interact with D3/MAX2 and D14 proteins (Wang et al. 2015; Soundappan et al. 2015). Interestingly, artificial miRNAs (amiRNAs) targeting SMXL6, SMXL7 and SMXL8 transcripts suppressed the max2-specific increase in shoot branching but not amiRNAs targeting the sub-clade 2 members SMXL4 and SMXL5 (Soundappan et al. 2015). Although the third sub-clade member, SMXL3, was not repressed in smxl45-ami max2 plants, these results are in agreement with the idea that members of clade 3 mediate SL signaling while other SMXL proteins fulfill different functions (Wang et al. 2015; Soundappan et al. 2015).

Supporting this assumption, the RGKTGI motif identified to be important for SL/KAR-dependent degradation (Jiang et al. 2013; Zhou et al. 2013; Soundappan et al. 2015) is not conserved in SMXL proteins belonging to clade 2 (Fig. 2). This opens up the possibility that members of this clade are SL/KAR-independent reminiscent to the situation in the GRAS family from which only a subset is GA-dependent. However, expression patterns of different family members are very diverse (Stanga et al. 2013; Soundappan et al. 2015) making it possible that, when compared to the DELLAs, the emerging differences in function are simply due to different sites of action. Looking again at GA signaling, posttranslational modification is important for DELLA activity. O-GlcNAcylation catalyzed by the GlcNAc transferase SPINDLY (SPY) promotes DELLA activity (Silverstone et al. 2007). Moreover, stress-dependent SUMOylation of DELLAs allows stable binding to GID1 independently from GA, resulting in reduced degradation of non-SUMOylated DELLAs and, therefore, decreased GA-sensitivity (Conti et al. 2014). Thus, presence or absence of SMXL proteins may not be the only critical aspect for determining the level of SL signaling in particular contexts.

The complexity of downstream processes

DELLAs, similarly to D53/SMXL proteins, do not contain a canonical DNA binding domain. However, DELLAs interact with several groups of transcription factors, thereby, preventing their DNA binding (Xu et al. 2014). Famous examples are the PHYTOCHROME INTERACTING FACTORS (PIFs). GA-dependent DELLA degradation releases these basic helix-loop-helix (bHLH) transcription factors and induces the transcription of genes which are conversely regulated by light through phytochrome-dependent PIF degradation (Huq and Quail 2002; Khanna et al. 2004; de Lucas et al. 2008; Feng et al. 2008). Thus, GA- and light signaling converge on the level of PIF transcription factors, nicely demonstrating how opposing stimuli are integrated on the molecular level. Likewise, DELLAs stimulate jasmonic acid (JA) signaling by titrating away JA ZIM-domain (JAZ) proteins acting as JA signaling repressors (Hou et al. 2010) and dampen brassinosteroid (BR) signaling by binding to the BRASSINAZOLE-RESISTANT1 (BZR1) transcription factor important for BR-dependent gene activation (Gallego-Bartolome et al. 2012; Bai et al. 2012). These findings reveal an astonishing broadness of direct interactive connections between different hormone-dependent transcriptional regulators and underline the necessity for integrative approaches to understand downstream responses.

In addition to interfering with the activity of other transcription factors, evidence for a direct stimulation of transcription has been documented, for example for SLR1 from rice (Hirano et al. 2012). The mystery of how DELLAs interact with DNA in this context has been elucidated recently by the identification of the DNA-binding INDETERMINATE DOMAIN (IDD) family proteins, which serve as transcriptional scaffolds in Arabidopsis (Yoshida et al. 2014). This study shows that IDD proteins are important for GA signaling and bind to both, the promoter of the SCARECROW-LIKE3 (SCL3) gene and to the RGA protein (Yoshida et al. 2014).

Beyond the direct or indirect regulation of transcription, DELLAs also titrate away proteins that move from the nucleus to the cytoplasm upon DELLA degradation to execute their function. In particular, the prefoldin complex (PFD), a co-chaperone required for tubulin folding, translocates after GA-induced DELLA degradation and increases the amount of active tubulin subunits promoting cell expansion (Locascio et al. 2013). Thus, DELLAs act as central hubs for executing GA signaling and integrating various signaling pathways on multiple cellular levels.

The molecular role of D53/SMXL proteins is still obscure. They are large (around 1000 amino acids) providing plenty of opportunities for interactions with other molecules. Indeed, D53/SMXL proteins carry a putative ethylene-responsive element binding factor-associated amphiphilic repression (EAR) domain that can interact with TOPLESS (TPL) (Jiang et al. 2013; Soundappan et al. 2015; Wang et al. 2015). TLP and TLP- RELATED (TRP) proteins are well studied repressors of transcription in plants and were found to specifically interact with transcription factors to regulate many growth processes (Causier et al. 2012). D53, SMAX1, SMXL6, SMXL7 and SMXL8 interact with TLP proteins in heterologous expression systems and in vitro (Jiang et al. 2013; Soundappan et al. 2015; Wang et al. 2015). Although the functional relevance of these interactions remains to be tested, this connection may help identifying downstream targets of SL-signaling and mechanisms of SL-dependent gene regulation.

Interestingly, SL signaling has been proposed to act in parallel to light perception by preventing the E3 ubiquitin-ligase CONSTITUTIVE PHOTOMORPHOGENIC1 (COP1) from entering the nucleus and degrading the light-responsive protein LONG HYPOCOTYL5 (HY5) (Tsuchiya et al. 2010; Jia et al. 2014). HY5 is a bZIP transcription factor antagonizing PIF activity by competing for the same promoter binding sites (Toledo-Ortiz et al. 2014). One of its best-known functions is the inhibition of hypocotyl elongation, which is used as a common readout to determine light- and/or SL/KAR-sensitivity in Arabidopsis (Jia et al. 2014; Scaffidi et al. 2013). GR24 suppresses hypocotyl elongation in a light- and MAX2-dependent manner (Jia et al. 2014). Moreover, hy5 and max2 mutants display an additive effect regarding GR24-insensitivity (Shen et al. 2012). Thus, although the exact molecular mechanism is so far unknown and highly debated, it has been hypothesized that MAX2 regulates photomorphogenesis (Jia et al. 2014; Waters and Smith 2013; Tsuchiya et al. 2010; Shen et al. 2012). However, as mentioned above, GR24 effects and a role of MAX2 are not necessarily indicative of a role of SL signaling in mediating the effect of light, as both are not specific for this pathway. Indeed, SL-deficient mutants usually do not display canonical light-related phenotypic alterations in a broad spectrum of species including Arabidopsis and pea (Urquhart et al. 2015; Shen et al. 2012). Furthermore, although hy5 and photoreceptor mutants are hyposensitive against GR24 and KAR treatments with respect to the repression of hypocotyl elongation (Jia et al. 2014; Waters and Smith 2013), molecular responses are not affected (Waters and Smith 2013) suggesting that SL-signaling, as such, is not part of the classic light signaling network.

Apart from being secreted by plant roots and their role in biotic interactions (Xie and Yoneyama 2010), SLs are best known as branching inhibitors (Brewer et al. 2009; Gomez-Roldan et al. 2008). In this case, a negative effect on polar auxin transport by reducing the amount of PIN- FORMED (PIN) auxin exporters in the plasma membrane has been demonstrated (Bennett et al. 2006; Shinohara et al. 2013). Computational modeling supports the idea that limiting auxin transport capacities is a crucial function of SLs in branching control. In this context, SLs enhance competition of branches for auxin transport capacities rather than acting as constitutive inhibitors (Crawford et al. 2010; Shinohara et al. 2013; Prusinkiewicz et al. 2009). In addition, local transcriptional activation of genes influencing branching, such as the TCP transcription factor BRANCHED1 (BRC1), has been described (Braun et al. 2012; Dun et al. 2012). Although the two roles of SL signaling in the regulation of branching has been discussed controversially (Brewer et al. 2015; Waldie et al. 2014), the multitude of direct targets of GA signaling, their parallel mode of action and spatial differences in the signaling process, provides a glimpse of the possible complexity and argues for an integration of different approaches.


Due to recent fundamental breakthroughs in SL biology research, we expect the unfolding of another complex signaling network in plants soon. In particular, the identification of the D53/SMXL protein family as repressors of SL signaling and direct targets of SL-dependent proteolysis opens up novel avenues to core events in the signaling cascade. Their characterization will be tremendously helpful for integrating the SL pathway into known regulatory networks and for understanding primary effects of SL signaling. Comparisons to other signaling cascades, like GA signaling, are certainly helpful as a first guideline in this regard. Such a comparison demonstrates the degree of complexity possible on the level of transport, perception, and targeted processes and emphasizes experimental pitfalls to be taken into consideration. For example, it will be essential to decipher the roles of the different SLs in planta and unwrap their distinct adaptive values. The spatio-temporal dynamics of SL signaling is another interesting aspect for which hardly any information is available. Do all cells have the capacity to sense SLs or is this mainly restricted to vascular tissues? Does sensitivity change over time or in different environments? The identification of events downstream of D53/SMXL proteolysis will certainly provide important insights and tools for addressing these questions. The dissimilarity of D53/SMXL proteins to any other group of known developmental regulators suggests the existence of unique molecular mechanisms and argues for surprising findings in the future.

Author contribution statement

EW, VL and TG wrote the manuscript. All authors read and approved the manuscript.



This work was supported by a Heisenberg Fellowship (GR 2104/3-1) and a research grant (GR 2104/4-1) from the German Research Foundation (DFG) and by an ERC Consolidator Grant (PLANTSTEMS, 647148). This article is based upon work from COST Action (FA1206), supported by COST (European Cooperation in Science and Technology). We thank Natalie Grace Schulz and Nial Rau Gursanscky for comments on the manuscript.


  1. Abe S, Sado A, Tanaka K, Kisugi T, Asami K, Ota S, Kim HI, Yoneyama K, Xie X, Ohnishi T, Seto Y, Yamaguchi S, Akiyama K, Yoneyama K, Nomura T (2014) Carlactone is converted to carlactonoic acid by MAX1 in Arabidopsis and its methyl ester can directly interact with AtD14 in vitro. Proc Natl Acad Sci USA 111(50):18084–18089PubMedPubMedCentralCrossRefGoogle Scholar
  2. Adamowski M, Friml J (2015) PIN-dependent auxin transport: action, regulation, and evolution. Plant Cell 27(1):20–32PubMedPubMedCentralCrossRefGoogle Scholar
  3. Agustí J, Herold S, Schwarz M, Sanchez P, Ljung K, Dun EA, Brewer PB, Beveridge CA, Sieberer T, Sehr EM, Greb T (2011) Strigolactone signaling is required for auxin-dependent stimulation of secondary growth in plants. Proc Natl Acad Sci USA 108(50):20242–20247PubMedPubMedCentralCrossRefGoogle Scholar
  4. Akiyama K, Matsuzaki K, Hayashi H (2005) Plant sesquiterpenes induce hyphal branching in arbuscular mycorrhizal fungi. Nature 435(7043):824–827PubMedCrossRefGoogle Scholar
  5. Akiyama K, Ogasawara S, Ito S, Hayashi H (2010) Structural requirements of strigolactones for hyphal branching in AM fungi. Plant Cell Physiol 51(7):1104–1117PubMedPubMedCentralCrossRefGoogle Scholar
  6. Ariizumi T, Murase K, Sun TP, Steber CM (2008) Proteolysis-independent downregulation of DELLA repression in Arabidopsis by the gibberellin receptor GIBBERELLIN INSENSITIVE DWARF1. Plant Cell 20(9):2447–2459PubMedPubMedCentralCrossRefGoogle Scholar
  7. Ariizumi T, Lawrence PK, Steber CM (2011) The role of two f-box proteins, SLEEPY1 and SNEEZY, in Arabidopsis gibberellin signaling. Plant Physiol 155(2):765–775PubMedPubMedCentralCrossRefGoogle Scholar
  8. Artuso E, Ghibaudi E, Lace B, Marabello D, Vinciguerra D, Lombardi C, Koltai H, Kapulnik Y, Novero M, Occhiato EG, Scarpi D, Parisotto S, Deagostino A, Venturello P, Mayzlish-Gati E, Bier A, Prandi C (2015) Stereochemical assignment of strigolactone analogues confirms their selective biological activity. J Nat Prod 78(11):2624–2633PubMedCrossRefGoogle Scholar
  9. Bai MY, Shang JX, Oh E, Fan M, Bai Y, Zentella R, Sun TP, Wang ZY (2012) Brassinosteroid, gibberellin and phytochrome impinge on a common transcription module in Arabidopsis. Nat Cell Biol 14(8):810–817PubMedPubMedCentralCrossRefGoogle Scholar
  10. Bennett T, Sieberer T, Willett B, Booker J, Luschnig C, Leyser O (2006) The Arabidopsis MAX pathway controls shoot branching by regulating auxin transport. Curr Biol 16(6):553–563PubMedCrossRefGoogle Scholar
  11. Bömke C, Tudzynski B (2009) Diversity, regulation, and evolution of the gibberellin biosynthetic pathway in fungi compared to plants and bacteria. Phytochemistry 70(15–16):1876–1893PubMedCrossRefGoogle Scholar
  12. Booker J, Sieberer T, Wright W, Williamson L, Willett B, Stirnberg P, Turnbull C, Srinivasan M, Goddard P, Leyser O (2005) MAX1 encodes a cytochrome P450 family member that acts downstream of MAX3/4 to produce a carotenoid-derived branch-inhibiting hormone. Dev Cell 8(3):443–449PubMedCrossRefGoogle Scholar
  13. Braun N, de Saint Germain A, Pillot JP, Boutet-Mercey S, Dalmais M, Antoniadi I, Li X, Maia-Grondard A, Le Signor C, Bouteiller N, Luo D, Bendahmane A, Turnbull C, Rameau C (2012) The pea TCP transcription factor PsBRC1 acts downstream of strigolactones to control shoot branching. Plant Physiol 158(1):225–238PubMedPubMedCentralCrossRefGoogle Scholar
  14. Brewer PB, Dun EA, Ferguson BJ, Rameau C, Beveridge CA (2009) Strigolactone acts downstream of auxin to regulate bud outgrowth in pea and Arabidopsis. Plant Physiol 150(1):482–493PubMedPubMedCentralCrossRefGoogle Scholar
  15. Brewer PB, Dun EA, Gui R, Mason MG, Beveridge CA (2015) Strigolactone inhibition of branching independent of polar auxin transport. Plant Physiol 168(4):1820–1829PubMedPubMedCentralCrossRefGoogle Scholar
  16. Cardoso C, Ruyter-Spira C, Bouwmeester HJ (2011) Strigolactones and root infestation by plant-parasitic Striga, Orobanche and Phelipanche spp. Plant Sci 180(3):414–420PubMedCrossRefGoogle Scholar
  17. Causier B, Lloyd J, Stevens L, Davies B (2012) TOPLESS co-repressor interactions and their evolutionary conservation in plants. Plant Signal Behav 7(3):325–328PubMedPubMedCentralCrossRefGoogle Scholar
  18. Chevalier F, Nieminen K, Sanchez-Ferrero JC, Rodriguez ML, Chagoyen M, Hardtke CS, Cubas P (2014) Strigolactone promotes degradation of DWARF14, an alpha/beta hydrolase essential for strigolactone signaling in Arabidopsis. Plant Cell 26(3):1134–1150PubMedPubMedCentralCrossRefGoogle Scholar
  19. Conn CE, Bythell-Douglas R, Neumann D, Yoshida S, Whittington B, Westwood JH, Shirasu K, Bond CS, Dyer KA, Nelson DC (2015) Convergent evolution of strigolactone perception enabled host detection in parasitic plants. Science 349(6247):540–543PubMedCrossRefGoogle Scholar
  20. Conti L, Nelis S, Zhang C, Woodcock A, Swarup R, Galbiati M, Tonelli C, Napier R, Hedden P, Bennett M, Sadanandom A (2014) Small ubiquitin-like modifier protein SUMO enables plants to control growth independently of the phytohormone gibberellin. Dev Cell 28(1):102–110PubMedCrossRefGoogle Scholar
  21. Cook CE, Whichard LP, Turner B, Wall ME, Egley GH (1966) Germination of witchweed (Striga lutea Lour.): isolation and properties of a potent stimulant. Science 154(3753):1189–1190PubMedCrossRefGoogle Scholar
  22. Crawford S, Shinohara N, Sieberer T, Williamson L, George G, Hepworth J, Muller D, Domagalska MA, Leyser O (2010) Strigolactones enhance competition between shoot branches by dampening auxin transport. Development 137(17):2905–2913PubMedCrossRefGoogle Scholar
  23. de Lucas M, Daviere JM, Rodriguez-Falcon M, Pontin M, Iglesias-Pedraz JM, Lorrain S, Fankhauser C, Blazquez MA, Titarenko E, Prat S (2008) A molecular framework for light and gibberellin control of cell elongation. Nature 451(7177):480–484PubMedCrossRefGoogle Scholar
  24. de Saint Germain A, Bonhomme S, Boyer FD, Rameau C (2013a) Novel insights into strigolactone distribution and signalling. Curr Opin Plant Biol 16(5):583–589CrossRefGoogle Scholar
  25. de Saint Germain A, Ligerot Y, Dun EA, Pillot JP, Ross JJ, Beveridge CA, Rameau C (2013b) Strigolactones stimulate internode elongation independently of gibberellins. Plant Physiol 163(2):1012–1025CrossRefGoogle Scholar
  26. Dill A, Thomas SG, Hu J, Steber CM, Sun TP (2004) The Arabidopsis F-box protein SLEEPY1 targets gibberellin signaling repressors for gibberellin-induced degradation. Plant Cell 16(6):1392–1405PubMedPubMedCentralCrossRefGoogle Scholar
  27. Domagalska MA, Leyser O (2011) Signal integration in the control of shoot branching. Nat Rev Mol Cell Biol 12(4):211–221PubMedCrossRefGoogle Scholar
  28. Dun EA, de Saint Germain A, Rameau C, Beveridge CA (2012) Antagonistic action of strigolactone and cytokinin in bud outgrowth control. Plant Physiol 158(1):487–498PubMedPubMedCentralCrossRefGoogle Scholar
  29. Eriksson S, Bohlenius H, Moritz T, Nilsson O (2006) GA4 is the active gibberellin in the regulation of LEAFY transcription and Arabidopsis floral initiation. Plant Cell 18(9):2172–2181PubMedPubMedCentralCrossRefGoogle Scholar
  30. Feng S, Martinez C, Gusmaroli G, Wang Y, Zhou J, Wang F, Chen L, Yu L, Iglesias-Pedraz JM, Kircher S, Schafer E, Fu X, Fan LM, Deng XW (2008) Coordinated regulation of Arabidopsis thaliana development by light and gibberellins. Nature 451(7177):475–479PubMedPubMedCentralCrossRefGoogle Scholar
  31. Foo E, Davies NW (2011) Strigolactones promote nodulation in pea. Planta 234(5):1073–1081PubMedCrossRefGoogle Scholar
  32. Fridlender M, Lace B, Wininger S, Dam A, Kumari P, Belausov E, Tsemach H, Kapulnik Y, Prandi C, Koltai H (2015) Influx and efflux of strigolactones are actively regulated and involve the cell-trafficking system. Mol Plant 8(12):1809–1812PubMedCrossRefGoogle Scholar
  33. Gallego-Bartolome J, Minguet EG, Marin JA, Prat S, Blazquez MA, Alabadi D (2010) Transcriptional diversification and functional conservation between DELLA proteins in Arabidopsis. Mol Biol Evol 27(6):1247–1256PubMedCrossRefGoogle Scholar
  34. Gallego-Bartolome J, Minguet EG, Grau-Enguix F, Abbas M, Locascio A, Thomas SG, Alabadi D, Blazquez MA (2012) Molecular mechanism for the interaction between gibberellin and brassinosteroid signaling pathways in Arabidopsis. Proc Natl Acad Sci USA 109(33):13446–13451PubMedPubMedCentralCrossRefGoogle Scholar
  35. Gomez-Roldan V, Fermas S, Brewer PB, Puech-Pages V, Dun EA, Pillot JP, Letisse F, Matusova R, Danoun S, Portais JC, Bouwmeester H, Becard G, Beveridge CA, Rameau C, Rochange SF (2008) Strigolactone inhibition of shoot branching. Nature 455(7210):189–194PubMedCrossRefGoogle Scholar
  36. Griffiths J, Murase K, Rieu I, Zentella R, Zhang ZL, Powers SJ, Gong F, Phillips AL, Hedden P, Sun TP, Thomas SG (2006) Genetic characterization and functional analysis of the GID1 gibberellin receptors in Arabidopsis. Plant Cell 18(12):3399–3414PubMedPubMedCentralCrossRefGoogle Scholar
  37. Guo Y, Zheng Z, La Clair JJ, Chory J, Noel JP (2013) Smoke-derived karrikin perception by the alpha/beta-hydrolase KAI2 from Arabidopsis. Proc Natl Acad Sci USA 110(20):8284–8289PubMedPubMedCentralCrossRefGoogle Scholar
  38. Gutjahr C (2014) Phytohormone signaling in arbuscular mycorhiza development. Curr Opin Plant Biol 20:26–34PubMedCrossRefGoogle Scholar
  39. Gutjahr C, Gobbato E, Choi J, Riemann M, Johnston MG, Summers W, Carbonnel S, Mansfield C, Yang S, Nadal M, Acosta I, Takano M, Jiao W, Schneeberger K, Kelly KA, Paszkowski U (2015) Rice perception of symbiotic arbuscular mycorrhizal fungi requires the karrikin receptor complex. Science 350(6267):1521–1524PubMedCrossRefGoogle Scholar
  40. Hamiaux C, Drummond RS, Janssen BJ, Ledger SE, Cooney JM, Newcomb RD, Snowden KC (2012) DAD2 is an alpha/beta hydrolase likely to be involved in the perception of the plant branching hormone, strigolactone. Curr Biol 22(21):2032–2036PubMedCrossRefGoogle Scholar
  41. Harberd NP, Belfield E, Yasumura Y (2009) The angiosperm gibberellin-GID1-DELLA growth regulatory mechanism: how an “inhibitor of an inhibitor” enables flexible response to fluctuating environments. Plant Cell 21(5):1328–1339PubMedPubMedCentralCrossRefGoogle Scholar
  42. Hauvermale AL, Ariizumi T, Steber CM (2014) The roles of the GA receptors GID1a, GID1b, and GID1c in sly1-independent GA signaling. Plant Signal Behav 9(2):e28030PubMedPubMedCentralCrossRefGoogle Scholar
  43. Hedden P, Thomas SG (2012) Gibberellin biosynthesis and its regulation. Biochem J 444(1):11–25PubMedCrossRefGoogle Scholar
  44. Hirano K, Kouketu E, Katoh H, Aya K, Ueguchi-Tanaka M, Matsuoka M (2012) The suppressive function of the rice DELLA protein SLR1 is dependent on its transcriptional activation activity. Plant J 71(3):443–453PubMedGoogle Scholar
  45. Hou X, Lee LY, Xia K, Yan Y, Yu H (2010) DELLAs modulate jasmonate signaling via competitive binding to JAZs. Dev Cell 19(6):884–894PubMedCrossRefGoogle Scholar
  46. Huq E, Quail PH (2002) PIF4, a phytochrome-interacting bHLH factor, functions as a negative regulator of phytochrome B signaling in Arabidopsis. EMBO J 21(10):2441–2450PubMedPubMedCentralCrossRefGoogle Scholar
  47. Iyer-Pascuzzi AS, Jackson T, Cui H, Petricka JJ, Busch W, Tsukagoshi H, Benfey PN (2011) Cell identity regulators link development and stress responses in the Arabidopsis root. Dev Cell 21(4):770–782PubMedPubMedCentralCrossRefGoogle Scholar
  48. Jia W, Davies WJ (2007) Modification of leaf apoplastic pH in relation to stomatal sensitivity to root-sourced abscisic acid signals. Plant Physiol 143(1):68–77PubMedPubMedCentralCrossRefGoogle Scholar
  49. Jia KP, Luo Q, He SB, Lu XD, Yang HQ (2014) Strigolactone-regulated hypocotyl elongation is dependent on cryptochrome and phytochrome signaling pathways in Arabidopsis. Mol Plant 7(3):528–540PubMedCrossRefGoogle Scholar
  50. Jiang L, Liu X, Xiong G, Liu H, Chen F, Wang L, Meng X, Liu G, Yu H, Yuan Y, Yi W, Zhao L, Ma H, He Y, Wu Z, Melcher K, Qian Q, Xu HE, Wang Y, Li J (2013) DWARF 53 acts as a repressor of strigolactone signalling in rice. Nature 504(7480):401–405PubMedCrossRefGoogle Scholar
  51. Kagiyama M, Hirano Y, Mori T, Kim SY, Kyozuka J, Seto Y, Yamaguchi S, Hakoshima T (2013) Structures of D14 and D14L in the strigolactone and karrikin signaling pathways. Genes Cells 18(2):147–160PubMedCrossRefGoogle Scholar
  52. Khanna R, Huq E, Kikis EA, Al-Sady B, Lanzatella C, Quail PH (2004) A novel molecular recognition motif necessary for targeting photoactivated phytochrome signaling to specific basic helix-loop-helix transcription factors. Plant Cell 16(11):3033–3044PubMedPubMedCentralCrossRefGoogle Scholar
  53. Kohlen W, Charnikhova T, Liu Q, Bours R, Domagalska MA, Beguerie S, Verstappen F, Leyser O, Bouwmeester HJ, Ruyter-Spira C (2011) Strigolactones are transported through the xylem and play a key role in shoot architectural response to phosphate deficiency in non-AM host Arabidopsis thaliana. Plant Physiol 155(2):974–987PubMedPubMedCentralCrossRefGoogle Scholar
  54. Kohlen W, Charnikhova T, Lammers M, Pollina T, Toth P, Haider I, Pozo MJ, de Maagd RA, Ruyter-Spira C, Bouwmeester HJ, Lopez-Raez JA (2012) The tomato CAROTENOID CLEAVAGE DIOXYGENASE8 (SlCCD8) regulates rhizosphere signaling, plant architecture and affects reproductive development through strigolactone biosynthesis. New Phytol 196(2):535–547PubMedCrossRefGoogle Scholar
  55. Kretzschmar T, Kohlen W, Sasse J, Borghi L, Schlegel M, Bachelier JB, Reinhardt D, Bours R, Bouwmeester HJ, Martinoia E (2012) A petunia ABC protein controls strigolactone-dependent symbiotic signalling and branching. Nature 483(7389):341–344PubMedCrossRefGoogle Scholar
  56. Larrieu A, Vernoux T (2015) Comparison of plant hormone signalling systems. Essays Biochem 58:165–181PubMedCrossRefGoogle Scholar
  57. Locascio A, Blazquez MA, Alabadi D (2013) Dynamic regulation of cortical microtubule organization through prefoldin-DELLA interaction. Curr Biol 23(9):804–809PubMedCrossRefGoogle Scholar
  58. MacMillan J (2001) Occurrence of gibberellins in vascular plants, fungi, and bacteria. J Plant Regul 20(4):387–442CrossRefGoogle Scholar
  59. Mangnus EM, Zwanenburg B (1992) Tentative molecular mechanism for germination stimulation of Striga and Orobanche seeds by strigol and its synthetic analogs. J Agric Food Chem 40(6):1066–1070CrossRefGoogle Scholar
  60. Nakajima M, Shimada A, Takashi Y, Kim YC, Park SH, Ueguchi-Tanaka M, Suzuki H, Katoh E, Iuchi S, Kobayashi M, Maeda T, Matsuoka M, Yamaguchi I (2006) Identification and characterization of Arabidopsis gibberellin receptors. Plant J 46(5):880–889PubMedCrossRefGoogle Scholar
  61. Nakamura H, Xue YL, Miyakawa T, Hou F, Qin HM, Fukui K, Shi X, Ito E, Ito S, Park SH, Miyauchi Y, Asano A, Totsuka N, Ueda T, Tanokura M, Asami T (2013) Molecular mechanism of strigolactone perception by DWARF14. Nat Commun 4:2613PubMedGoogle Scholar
  62. Nelson DC, Scaffidi A, Dun EA, Waters MT, Flematti GR, Dixon KW, Beveridge CA, Ghisalberti EL, Smith SM (2011) F-box protein MAX2 has dual roles in karrikin and strigolactone signaling in Arabidopsis thaliana. Proc Natl Acad Sci USA 108(21):8897–8902PubMedPubMedCentralCrossRefGoogle Scholar
  63. Ni J, Gao C, Chen MS, Pan BZ, Ye K, Xu ZF (2015) Gibberellin promotes shoot branching in the perennial woody plant Jatropha curcas. Plant Cell Physiol 56(8):1655–1666PubMedPubMedCentralCrossRefGoogle Scholar
  64. Nir I, Moshelion M, Weiss D (2014) The Arabidopsis gibberellin methyl transferase 1 suppresses gibberellin activity, reduces whole-plant transpiration and promotes drought tolerance in transgenic tomato. Plant, Cell Environ 37(1):113–123CrossRefGoogle Scholar
  65. Nomura S, Nakashima H, Mizutani M, Takikawa H, Sugimoto Y (2013) Structural requirements of strigolactones for germination induction and inhibition of Striga gesnerioides seeds. Plant Cell Rep 32(6):829–838PubMedCrossRefGoogle Scholar
  66. Prandi C, Rosso H, Lace B, Occhiato EG, Oppedisano A, Tabasso S, Alberto G, Blangetti M (2013) Strigolactone analogs as molecular probes in chasing the (SLs) receptor/s: design and synthesis of fluorescent labeled molecules. Mol Plant 6(1):113–127PubMedCrossRefGoogle Scholar
  67. Proebsting WM, Hedden P, Lewis MJ, Croker SJ, Proebsting LN (1992) Gibberellin concentration and transport in genetic lines of pea: effects of grafting. Plant Physiol 100(3):1354–1360PubMedPubMedCentralCrossRefGoogle Scholar
  68. Prusinkiewicz P, Crawford S, Smith RS, Ljung K, Bennett T, Ongaro V, Leyser O (2009) Control of bud activation by an auxin transport switch. Proc Natl Acad Sci USA 106(41):17431–17436PubMedPubMedCentralCrossRefGoogle Scholar
  69. Ragni L, Nieminen K, Pacheco-Villalobos D, Sibout R, Schwechheimer C, Hardtke CS (2011) Mobile gibberellin directly stimulates Arabidopsis hypocotyl xylem expansion. Plant Cell 23(4):1322–1336PubMedPubMedCentralCrossRefGoogle Scholar
  70. Rasmussen A, Depuydt S, Goormachtig S, Geelen D (2013a) Strigolactones fine-tune the root system. Planta 238(4):615–626PubMedCrossRefGoogle Scholar
  71. Rasmussen A, Heugebaert T, Matthys C, Van Deun R, Boyer FD, Goormachtig S, Stevens C, Geelen D (2013b) A fluorescent alternative to the synthetic strigolactone GR24. Mol Plant 6(1):100–112PubMedCrossRefGoogle Scholar
  72. Regnault T, Davière JM, Wild M, Sakvarelidze-Achard L, Heintz D, Carrera Bergua E, Lopez Diaz I, Gong F, Hedden P, Achard P (2015) The gibberellin precursor GA12 acts as a long-distance growth signal in Arabidopsis. Nat Plants 1:15073CrossRefGoogle Scholar
  73. Ruyter-Spira C, Kohlen W, Charnikhova T, van Zeijl A, van Bezouwen L, de Ruijter N, Cardoso C, Lopez-Raez JA, Matusova R, Bours R, Verstappen F, Bouwmeester H (2011) Physiological effects of the synthetic strigolactone analog GR24 on root system architecture in Arabidopsis: another belowground role for strigolactones? Plant Physiol 155(2):721–734PubMedPubMedCentralCrossRefGoogle Scholar
  74. Sasse J, Simon S, Gubeli C, Liu GW, Cheng X, Friml J, Bouwmeester H, Martinoia E, Borghi L (2015) Asymmetric localizations of the ABC transporter PaPDR1 trace paths of directional strigolactone transport. Curr Biol 25(5):647–655PubMedCrossRefGoogle Scholar
  75. Savaldi-Goldstein S, Peto C, Chory J (2007) The epidermis both drives and restricts plant shoot growth. Nature 446(7132):199–202PubMedCrossRefGoogle Scholar
  76. Scaffidi A, Waters MT, Ghisalberti EL, Dixon KW, Flematti GR, Smith SM (2013) Carlactone-independent seedling morphogenesis in Arabidopsis. Plant J 76(1):1–9PubMedGoogle Scholar
  77. Scaffidi A, Waters MT, Sun YK, Skelton BW, Dixon KW, Ghisalberti EL, Flematti GR, Smith SM (2014) Strigolactone hormones and their stereoisomers signal through two related receptor proteins to induce different physiological responses in Arabidopsis. Plant Physiol 165(3):1221–1232PubMedPubMedCentralCrossRefGoogle Scholar
  78. Schwechheimer C, Willige BC (2009) Shedding light on gibberellic acid signalling. Curr Opin Plant Biol 12(1):57–62PubMedCrossRefGoogle Scholar
  79. Seto Y, Yamaguchi S (2014) Strigolactone biosynthesis and perception. Curr Opin Plant Biol 21:1–6PubMedCrossRefGoogle Scholar
  80. Shani E, Weinstain R, Zhang Y, Castillejo C, Kaiserli E, Chory J, Tsien RY, Estelle M (2013) Gibberellins accumulate in the elongating endodermal cells of Arabidopsis root. Proc Natl Acad Sci USA 110(12):4834–4839PubMedPubMedCentralCrossRefGoogle Scholar
  81. Shen H, Zhu L, Bu QY, Huq E (2012) MAX2 affects multiple hormones to promote photomorphogenesis. Mol Plant 5(3):750–762PubMedCrossRefGoogle Scholar
  82. Shimada A, Ueguchi-Tanaka M, Nakatsu T, Nakajima M, Naoe Y, Ohmiya H, Kato H, Matsuoka M (2008) Structural basis for gibberellin recognition by its receptor GID1. Nature 456(7221):520–523PubMedCrossRefGoogle Scholar
  83. Shinohara N, Taylor C, Leyser O (2013) Strigolactone can promote or inhibit shoot branching by triggering rapid depletion of the auxin efflux protein PIN1 from the plasma membrane. PLoS Biol 11(1):e1001474PubMedPubMedCentralCrossRefGoogle Scholar
  84. Silverstone AL, Tseng TS, Swain SM, Dill A, Jeong SY, Olszewski NE, Sun TP (2007) Functional analysis of SPINDLY in gibberellin signaling in Arabidopsis. Plant Physiol 143(2):987–1000PubMedPubMedCentralCrossRefGoogle Scholar
  85. Soundappan I, Bennett T, Morffy N, Liang Y, Stanga JP, Abbas A, Leyser O, Nelson DC (2015) SMAX1-LIKE/D53 family members enable distinct MAX2-dependent responses to strigolactones and karrikins in Arabidopsis. Plant Cell 27(11):3143–3159PubMedCrossRefGoogle Scholar
  86. Stanga JP, Smith SM, Briggs WR, Nelson DC (2013) SUPPRESSOR OF MORE AXILLARY GROWTH2 1 controls seed germination and seedling development in Arabidopsis. Plant Physiol 163(1):318–330PubMedPubMedCentralCrossRefGoogle Scholar
  87. Stanga JP, Morffy N, Nelson DC (2016) Functional redundancy in the control of seedling growth by the karrikin signaling pathway. Planta. doi: 10.1007/s00425-015-2458-2 PubMedGoogle Scholar
  88. Stirnberg P, Furner IJ, Leyser HMO (2007) MAX2 participates in an SCF complex which acts locally at the node to suppress shoot branching. Plant J 50(1):80–94PubMedCrossRefGoogle Scholar
  89. Sun TP (2011) The molecular mechanism and evolution of the GA-GID1-DELLA signaling module in plants. Curr Biol 21(9):R338–R345PubMedCrossRefGoogle Scholar
  90. Talon M, Koornneef M, Zeevaart JA (1990) Endogenous gibberellins in Arabidopsis thaliana and possible steps blocked in the biosynthetic pathways of the semidwarf ga4 and ga5 mutants. Proc Natl Acad Sci USA 87(20):7983–7987PubMedPubMedCentralCrossRefGoogle Scholar
  91. Toh S, Holbrook-Smith D, Stogios PJ, Onopriyenko O, Lumba S, Tsuchiya Y, Savchenko A, McCourt P (2015) Structure-function analysis identifies highly sensitive strigolactone receptors in Striga. Science 350(6257):203–207PubMedCrossRefGoogle Scholar
  92. Toledo-Ortiz G, Johansson H, Lee KP, Bou-Torrent J, Stewart K, Steel G, Rodriguez-Concepcion M, Halliday KJ (2014) The HY5-PIF regulatory module coordinates light and temperature control of photosynthetic gene transcription. PLoS Genet 10(6):e1004416PubMedPubMedCentralCrossRefGoogle Scholar
  93. Tsuchiya Y, Vidaurre D, Toh S, Hanada A, Nambara E, Kamiya Y, Yamaguchi S, McCourt P (2010) A small-molecule screen identifies new functions for the plant hormone strigolactone. Nat Chem Biol 6(10):741–749PubMedCrossRefGoogle Scholar
  94. Turnbull CG, Booker JP, Leyser HM (2002) Micrografting techniques for testing long-distance signalling in Arabidopsis. Plant J 32(2):255–262PubMedCrossRefGoogle Scholar
  95. Ubeda-Tomas S, Swarup R, Coates J, Swarup K, Laplaze L, Beemster GT, Hedden P, Bhalerao R, Bennett MJ (2008) Root growth in Arabidopsis requires gibberellin/DELLA signalling in the endodermis. Nat Cell Biol 10(5):625–628PubMedCrossRefGoogle Scholar
  96. Ueguchi-Tanaka M, Matsuoka M (2010) The perception of gibberellins: clues from receptor structure. Curr Opin Plant Biol 13(5):503–508PubMedCrossRefGoogle Scholar
  97. Ueguchi-Tanaka M, Ashikari M, Nakajima M, Itoh H, Katoh E, Kobayashi M, Chow TY, Hsing YI, Kitano H, Yamaguchi I, Matsuoka M (2005) GIBBERELLIN INSENSITIVE DWARF1 encodes a soluble receptor for gibberellin. Nature 437(7059):693–698PubMedCrossRefGoogle Scholar
  98. Ueguchi-Tanaka M, Nakajima M, Motoyuki A, Matsuoka M (2007) Gibberellin receptor and its role in gibberellin signaling in plants. Annu Rev Plant Biol 58:183–198PubMedCrossRefGoogle Scholar
  99. Ueguchi-Tanaka M, Hirano K, Hasegawa Y, Kitano H, Matsuoka M (2008) Release of the repressive activity of rice DELLA protein SLR1 by gibberellin does not require SLR1 degradation in the gid2 mutant. Plant Cell 20(9):2437–2446PubMedPubMedCentralCrossRefGoogle Scholar
  100. Ueno K, Furumoto T, Umeda S, Mizutani M, Takikawa H, Batchvarova R, Sugimoto Y (2014) Heliolactone, a non-sesquiterpene lactone germination stimulant for root parasitic weeds from sunflower. Phytochemistry 108:122–128PubMedCrossRefGoogle Scholar
  101. Umehara M, Hanada A, Yoshida S, Akiyama K, Arite T, Takeda-Kamiya N, Magome H, Kamiya Y, Shirasu K, Yoneyama K, Kyozuka J, Yamaguchi S (2008) Inhibition of shoot branching by new terpenoid plant hormones. Nature 455(7210):195–200PubMedCrossRefGoogle Scholar
  102. Umehara M, Hanada A, Magome H, Takeda-Kamiya N, Yamaguchi S (2010) Contribution of strigolactones to the inhibition of tiller bud outgrowth under phosphate deficiency in rice. Plant Cell Physiol 51(7):1118–1126PubMedPubMedCentralCrossRefGoogle Scholar
  103. Umehara M, Cao M, Akiyama K, Akatsu T, Seto Y, Hanada A, Li W, Takeda-Kamiya N, Morimoto Y, Yamaguchi S (2015) Structural requirements of strigolactones for shoot branching inhibition in rice and Arabidopsis. Plant Cell Physiol 56(6):1059–1072PubMedCrossRefGoogle Scholar
  104. Urquhart S, Foo E, Reid JB (2015) The role of strigolactones in photomorphogenesis of pea is limited to adventitious rooting. Physiol Plant 153(3):392–402PubMedCrossRefGoogle Scholar
  105. Varbanova M, Yamaguchi S, Yang Y, McKelvey K, Hanada A, Borochov R, Yu F, Jikumaru Y, Ross J, Cortes D, Ma CJ, Noel JP, Mander L, Shulaev V, Kamiya Y, Rodermel S, Weiss D, Pichersky E (2007) Methylation of gibberellins by Arabidopsis GAMT1 and GAMT2. Plant Cell 19(1):32–45PubMedPubMedCentralCrossRefGoogle Scholar
  106. Waldie T, McCulloch H, Leyser O (2014) Strigolactones and the control of plant development: lessons from shoot branching. Plant J 79(4):607–622PubMedCrossRefGoogle Scholar
  107. Wang F, Deng XW (2011) Plant ubiquitin-proteasome pathway and its role in gibberellin signaling. Cell Res 21(9):1286–1294PubMedPubMedCentralCrossRefGoogle Scholar
  108. Wang F, Zhu D, Huang X, Li S, Gong Y, Yao Q, Fu X, Fan LM, Deng XW (2009) Biochemical insights on degradation of Arabidopsis DELLA proteins gained from a cell-free assay system. Plant Cell 21(8):2378–2390PubMedPubMedCentralCrossRefGoogle Scholar
  109. Wang L, Wang B, Jiang L, Liu X, Li X, Lu Z, Meng X, Wang Y, Smith SM, Li J (2015) Strigolactone signaling in Arabidopsis regulates shoot development by targeting D53-Like SMXL repressor proteins for ubiquitination and degradation. Plant Cell 27(11):3128–3142PubMedPubMedCentralCrossRefGoogle Scholar
  110. Waters MT, Smith SM (2013) KAI2- and MAX2-mediated responses to karrikins and strigolactones are largely independent of HY5 in Arabidopsis seedlings. Mol Plant 6(1):63–75PubMedCrossRefGoogle Scholar
  111. Waters MT, Nelson DC, Scaffidi A, Flematti GR, Sun YK, Dixon KW, Smith SM (2012) Specialisation within the DWARF14 protein family confers distinct responses to karrikins and strigolactones in Arabidopsis. Development 139(7):1285–1295PubMedCrossRefGoogle Scholar
  112. Waters MT, Scaffidi A, Sun YK, Flematti GR, Smith SM (2014) The karrikin response system of Arabidopsis. Plant J 79(4):623–631PubMedCrossRefGoogle Scholar
  113. Xie X, Yoneyama K (2010) The strigolactone story. Annu Rev Phytopathol 48:93–117PubMedCrossRefGoogle Scholar
  114. Xu H, Liu Q, Yao T, Fu X (2014) Shedding light on integrative GA signaling. Curr Opin Plant Biol 21:89–95PubMedCrossRefGoogle Scholar
  115. Yoshida H, Hirano K, Sato T, Mitsuda N, Nomoto M, Maeo K, Koketsu E, Mitani R, Kawamura M, Ishiguro S, Tada Y, Ohme-Takagi M, Matsuoka M, Ueguchi-Tanaka M (2014) DELLA protein functions as a transcriptional activator through the DNA binding of the indeterminate domain family proteins. Proc Natl Acad Sci USA 111(21):7861–7866PubMedPubMedCentralCrossRefGoogle Scholar
  116. Zhang ZL, Ogawa M, Fleet CM, Zentella R, Hu J, Heo JO, Lim J, Kamiya Y, Yamaguchi S, Sun TP (2011) Scarecrow-like 3 promotes gibberellin signaling by antagonizing master growth repressor DELLA in Arabidopsis. Proc Natl Acad Sci USA 108(5):2160–2165PubMedPubMedCentralCrossRefGoogle Scholar
  117. Zhang Y, van Dijk AD, Scaffidi A, Flematti GR, Hofmann M, Charnikhova T, Verstappen F, Hepworth J, van der Krol S, Leyser O, Smith SM, Zwanenburg B, Al-Babili S, Ruyter-Spira C, Bouwmeester HJ (2014) Rice cytochrome P450 MAX1 homologs catalyze distinct steps in strigolactone biosynthesis. Nat Chem Biol 10(12):1028–1033PubMedCrossRefGoogle Scholar
  118. Zhao LH, Zhou XE, Yi W, Wu Z, Liu Y, Kang Y, Hou L, de Waal PW, Li S, Jiang Y, Scaffidi A, Flematti GR, Smith SM, Lam VQ, Griffin PR, Wang Y, Li J, Melcher K, Xu HE (2015) Destabilization of strigolactone receptor DWARF14 by binding of ligand and E3-ligase signaling effector DWARF3. Cell Res 25(11):1219–1236PubMedCrossRefGoogle Scholar
  119. Zhou F, Lin Q, Zhu L, Ren Y, Zhou K, Shabek N, Wu F, Mao H, Dong W, Gan L, Ma W, Gao H, Chen J, Yang C, Wang D, Tan J, Zhang X, Guo X, Wang J, Jiang L, Liu X, Chen W, Chu J, Yan C, Ueno K, Ito S, Asami T, Cheng Z, Lei C, Zhai H, Wu C, Wang H, Zheng N, Wan J (2013) D14-SCF(D3)-dependent degradation of D53 regulates strigolactone signalling. Nature 504(7480):406–410PubMedPubMedCentralCrossRefGoogle Scholar
  120. Zhu Y, Nomura T, Xu Y, Zhang Y, Peng Y, Mao B, Hanada A, Zhou H, Wang R, Li P, Zhu X, Mander LN, Kamiya Y, Yamaguchi S, He Z (2006) ELONGATED UPPERMOST INTERNODE encodes a cytochrome P450 monooxygenase that epoxidizes gibberellins in a novel deactivation reaction in rice. Plant Cell 18(2):442–456PubMedPubMedCentralCrossRefGoogle Scholar
  121. Zwanenburg B, Pospisil T (2013) Structure and activity of strigolactones: new plant hormones with a rich future. Mol Plant 6(1):38–62PubMedCrossRefGoogle Scholar
  122. Zwanenburg B, Nayak SK, Charnikhova TV, Bouwmeester HJ (2013) New strigolactone mimics: structure-activity relationship and mode of action as germinating stimulants for parasitic weeds. Bioorg Med Chem Lett 23(18):5182–5186PubMedCrossRefGoogle Scholar
  123. Zwanenburg B, Zeljkovic SC, Pospisil T (2015) Synthesis of strigolactones, a strategic account. Pest Manag Sci. doi: 10.1002/ps.4105 PubMedGoogle Scholar

Copyright information

© The Author(s) 2016

Open AccessThis article is distributed under the terms of the Creative Commons Attribution 4.0 International License (, which permits unrestricted use, distribution, and reproduction in any medium, provided you give appropriate credit to the original author(s) and the source, provide a link to the Creative Commons license, and indicate if changes were made.

Authors and Affiliations

  • Eva-Sophie Wallner
    • 1
  • Vadir López-Salmerón
    • 1
  • Thomas Greb
    • 1
    Email author
  1. 1.Centre for Organismal Studies (COS)Heidelberg UniversityHeidelbergGermany

Personalised recommendations