In Vitro Dynamic and Quantitative Monitoring of Strigolactone-signaling Complex Formation by Time-resolved FRET

In Vitro Dynamic and Quantitative Monitoring of Strigolactone-signaling Complex Formation by Time-resolved FRET | bioRxiv /* */ /* */ <!-- <!-- /*! * yepnope1.5.4 * (c) WTFPL, GPLv2 */ (function(a,b,c){function d(a){return"[object Function]"==o.call(a)}function e(a){return"string"==typeof a}function f(){}function g(a){return!a||"loaded"==a||"complete"==a||"uninitialized"==a}function h(){var a=p.shift();q=1,a?a.t?m(function(){("c"==a.t?B.injectCss:B.injectJs)(a.s,0,a.a,a.x,a.e,1)},0):(a(),h()):q=0}function i(a,c,d,e,f,i,j){function k(b){if(!o&&g(l.readyState)&&(u.r=o=1,!q&&h(),l.onload=l.onreadystatechange=null,b)){"img"!=a&&m(function(){t.removeChild(l)},50);for(var d in y[c])y[c].hasOwnProperty(d)&&y[c][d].onload()}}var j=j||B.errorTimeout,l=b.createElement(a),o=0,r=0,u={t:d,s:c,e:f,a:i,x:j};1===y[c]&&(r=1,y[c]=[]),"object"==a?l.data=c:(l.src=c,l.type=a),l.width=l.height="0",l.onerror=l.onload=l.onreadystatechange=function(){k.call(this,r)},p.splice(e,0,u),"img"!=a&&(r||2===y[c]?(t.insertBefore(l,s?null:n),m(k,j)):y[c].push(l))}function j(a,b,c,d,f){return q=0,b=b||"j",e(a)?i("c"==b?v:u,a,b,this.i++,c,d,f):(p.splice(this.i++,0,a),1==p.length&&h()),this}function k(){var a=B;return a.loader={load:j,i:0},a}var l=b.documentElement,m=a.setTimeout,n=b.getElementsByTagName("script")[0],o={}.toString,p=[],q=0,r="MozAppearance"in l.style,s=r&&!!b.createRange().compareNode,t=s?l:n.parentNode,l=a.opera&&"[object Opera]"==o.call(a.opera),l=!!b.attachEvent&&!l,u=r?"object":l?"script":"img",v=l?"script":u,w=Array.isArray||function(a){return"[object Array]"==o.call(a)},x=[],y={},z={timeout:function(a,b){return b.length&&(a.timeout=b[0]),a}},A,B;B=function(a){function b(a){var a=a.split("!"),b=x.length,c=a.pop(),d=a.length,c={url:c,origUrl:c,prefixes:a},e,f,g;for(f=0;f<d;f++)g=a[f].split("="),(e=z[g.shift()])&&(c=e(c,g));for(f=0;f<b;f++)c=x[f](c);return c}function g(a,e,f,g,h){var i=b(a),j=i.autoCallback;i.url.split(".").pop().split("?").shift(),i.bypass||(e&&(e=d(e)?e:e[a]||e[g]||e[a.split("/").pop().split("?")[0]]),i.instead?i.instead(a,e,f,g,h):(y[i.url]?i.noexec=!0:y[i.url]=1,f.load(i.url,i.forceCSS||!i.forceJS&&"css"==i.url.split(".").pop().split("?").shift()?"c":c,i.noexec,i.attrs,i.timeout),(d(e)||d(j))&&f.load(function(){k(),e&&e(i.origUrl,h,g),j&&j(i.origUrl,h,g),y[i.url]=2})))}function h(a,b){function c(a,c){if(a){if(e(a))c||(j=function(){var a=[].slice.call(arguments);k.apply(this,a),l()}),g(a,j,b,0,h);else if(Object(a)===a)for(n in m=function(){var b=0,c;for(c in a)a.hasOwnProperty(c)&&b++;return b}(),a)a.hasOwnProperty(n)&&(!c&&!--m&&(d(j)?j=function(){var a=[].slice.call(arguments);k.apply(this,a),l()}:j[n]=function(a){return function(){var b=[].slice.call(arguments);a&&a.apply(this,b),l()}}(k[n])),g(a[n],j,b,n,h))}else!c&&l()}var h=!!a.test,i=a.load||a.both,j=a.callback||f,k=j,l=a.complete||f,m,n;c(h?a.yep:a.nope,!!i),i&&c(i)}var i,j,l=this.yepnope.loader;if(e(a))g(a,0,l,0);else if(w(a))for(i=0;i (function(w,d,s,l,i){w[l]=w[l]||[];w[l].push({'gtm.start':new Date().getTime(),event:'gtm.js'});var f=d.getElementsByTagName(s)[0];var j=d.createElement(s);var dl=l!='dataLayer'?'&l='+l:'';j.src='//www.googletagmanager.com/gtm.js?id='+i+dl;j.type='text/javascript';j.async=true;f.parentNode.insertBefore(j,f);})(window,document,'script','dataLayer','GTM-M677548'); Skip to main content Home About Submit ALERTS / RSS Search for this keyword Advanced Search New Results In Vitro Dynamic and Quantitative Monitoring of Strigolactone-signaling Complex Formation by Time-resolved FRET Taiki Suzuki , Kotaro Nishiyama , Yusuke Kato , Chihiro Shinkai , Tomoya Ishikawa , Jekson Robertlee , Michio Kuruma , Shinya Hagihara , Marco Bürger , Kosuke Fukui , Tadao Asami , Yoshiya Seto doi: https://doi.org/10.1101/2025.11.18.688998 Taiki Suzuki 1 Laboratory of Plant Chemical Regulation, School of Agriculture, Meiji University , 1-1-1 Higashi-Mita, Tama-ku, Kawasaki, Kanagawa 214-8571, Japan Find this author on Google Scholar Find this author on PubMed Search for this author on this site Kotaro Nishiyama 1 Laboratory of Plant Chemical Regulation, School of Agriculture, Meiji University , 1-1-1 Higashi-Mita, Tama-ku, Kawasaki, Kanagawa 214-8571, Japan Find this author on Google Scholar Find this author on PubMed Search for this author on this site For correspondence: nishiyama{at}meiji.ac.jp yoshiya{at}meiji.ac.jp Yusuke Kato 1 Laboratory of Plant Chemical Regulation, School of Agriculture, Meiji University , 1-1-1 Higashi-Mita, Tama-ku, Kawasaki, Kanagawa 214-8571, Japan Find this author on Google Scholar Find this author on PubMed Search for this author on this site Chihiro Shinkai 1 Laboratory of Plant Chemical Regulation, School of Agriculture, Meiji University , 1-1-1 Higashi-Mita, Tama-ku, Kawasaki, Kanagawa 214-8571, Japan Find this author on Google Scholar Find this author on PubMed Search for this author on this site Tomoya Ishikawa 1 Laboratory of Plant Chemical Regulation, School of Agriculture, Meiji University , 1-1-1 Higashi-Mita, Tama-ku, Kawasaki, Kanagawa 214-8571, Japan Find this author on Google Scholar Find this author on PubMed Search for this author on this site Jekson Robertlee 2 RIKEN Center for Sustainable Resource Science , 2-1 Hirosawa, Wako, Saitama 351-0198, Japan Find this author on Google Scholar Find this author on PubMed Search for this author on this site Michio Kuruma 1 Laboratory of Plant Chemical Regulation, School of Agriculture, Meiji University , 1-1-1 Higashi-Mita, Tama-ku, Kawasaki, Kanagawa 214-8571, Japan 2 RIKEN Center for Sustainable Resource Science , 2-1 Hirosawa, Wako, Saitama 351-0198, Japan Find this author on Google Scholar Find this author on PubMed Search for this author on this site Shinya Hagihara 2 RIKEN Center for Sustainable Resource Science , 2-1 Hirosawa, Wako, Saitama 351-0198, Japan Find this author on Google Scholar Find this author on PubMed Search for this author on this site Marco Bürger 3 Plant Biology Laboratory, Salk Institute for Biological Studies , 10010 North Torrey Pines Road, La Jolla, CA 92037, USA Find this author on Google Scholar Find this author on PubMed Search for this author on this site Kosuke Fukui 4 Faculty of Science Division I, Tokyo University of science , 1-3 Kagurazaka, Shinjuku-ku, Tokyo 162-8601, Japan Find this author on Google Scholar Find this author on PubMed Search for this author on this site Tadao Asami 5 Graduate School of Agricultural and Life Sciences, The University of Tokyo , 1-1-1 Yayoi, Bunkyo-ku, Tokyo 113-8657, Japan 6 Kihara Biological Institute, Yokohama City University , Maioka, Totsuka, Yokohama, Kanagawa 244-0813, Japan Find this author on Google Scholar Find this author on PubMed Search for this author on this site Yoshiya Seto 1 Laboratory of Plant Chemical Regulation, School of Agriculture, Meiji University , 1-1-1 Higashi-Mita, Tama-ku, Kawasaki, Kanagawa 214-8571, Japan Find this author on Google Scholar Find this author on PubMed Search for this author on this site For correspondence: nishiyama{at}meiji.ac.jp yoshiya{at}meiji.ac.jp Abstract Full Text Info/History Metrics Supplementary material Preview PDF Summary Strigolactones (SLs) are a class of plant hormones that play a critical role in the suppression of shoot branching. Furthermore, they are exuded from roots and act as signaling molecules for inter-organism communication in the rhizosphere. Strigolactones trigger those responses by inducing protein–protein interactions (PPIs) of signaling components and subsequent proteolysis of transcriptional repressors. The sequential event involves SL hydrolysis mediated by SL receptors belonging to an α/μ-hydrolase family, although the physiological role of SL hydrolysis is a subject of debate. To date, SL-induced PPIs have been analyzed by methods such as yeast-two hybrid, pull-down, and AlphaScreen assays. However, the kinetic aspect of PPI profiles has not been well studied. Here, we developed an in vitro method to monitor the formation of the SL signaling complex based on Time-Resolved Förster Resonance Energy Transfer (TR-FRET) technology. Our TR-FRET-based assay system allows us to analyze the mode of action of SL analogs from kinetic and quantitative perspectives. Notably, our method revealed differences in the intensity and time-dependency of PPI signals among different SL analogs with a range of hydrolyzabilities. In addition, we found that tolfenamic acid, an antagonist of the SL receptor, inhibited the SL-induced PPI but could not disrupt the already-formed signaling complex. The TR-FRET system was also used to rapidly and specifically detect naturally occurring SLs from root exudates containing many impurities. This work provides insights into the molecular mechanism of SL perception as well as a powerful tool for activity-based screening of SL signaling modulators. Significance statement The dual roles of strigolactone (SL) receptors in both the perception and deactivation of SLs make it difficult to elucidate the underlying molecular mechanism of SL signaling. We developed a new in vitro method to evaluate the dynamic activation of the SL receptor, and used it to gain deeper insights into the molecular mechanism of SL-signaling complex formation in response to the SL receptor agonists. Introduction Plant hormones regulate every aspect of plant life including growth, development, and responses to environmental stresses ( Santner et al ., 2009 ). Most plant hormones promote protein–protein interactions (PPIs) between receptors and signaling components ( Takeuchi et al ., 2021 ). The PPIs induce each downstream signal through protein phosphorylation or degradation. Thus, the development of assay methods to monitor hormone-induced PPIs, the key step in plant hormone signaling, is critical for thoroughly understanding the hormone signaling mechanism. Strigolactones (SLs) are the most recently identified class of plant hormones. They play an important role in the inhibition of shoot branching in angiosperms ( Umehara et al ., 2008 ; Gomez-Roldan et al ., 2008 ). In Arabidopsis thaliana , the SL receptor AtDWARF14 (AtD14) recognizes SL and subsequently recruits the transcriptional repressor proteins SUPPRESSOR OF MAX2 1-LIKE 6, 7, and 8 (SMXL6/7/8) and the F-box protein MORE AXIRALLY GROWTH 2 (MAX2), which is a component of the SCF E3 ubiquitin ligase complex, via physical interactions. After the ubiquitination of the repressor proteins, they are degraded through the 26S proteasome system, leading to the inhibition of shoot branching ( Wang et al ., 2015 ; Soundappan et al ., 2015 ; Yao et al ., 2016 ; Zhao et al ., 2015 ; Zhou et al ., 2013 ; Jiang et al ., 2013 ). Various SLs also play important roles in the rhizosphere as symbiotic signals for arbuscular mycorrhizal fungi ( Akiyama et al ., 2005 ). The SLs exuded from roots are exploited as germination stimulants by obligate root parasitic plants in Orobanchaceae family, which pose a serious threat to global food security ( Cook et al ., 1966 ). Proteins in the divergent clade KARRIKIN INSENSITIVE2/HYPOSENSITIVE TO LIGHT (KAI2/HTL), which is a paralogous family of AtD14, hereinafter called KAI2d, were identified as the receptors for host root-derived SLs in the obligate root parasitic plant Striga hermonthica ( Tsuchiya et al ., 2015 ; Toh et al ., 2015 ; Conn et al ., 2015 ). More recently, highly sensitive SL receptors were isolated and characterized from another obligate parasitic plant, Orobanche minor , and from the facultative hemiparasitic plant, Phtheirospermum japonicum ( Takei et al ., 2023 ; Takei et al ., 2024 ). Similar to AtD14, SL-dependent activation of KAI2d promotes PPIs among KAI2d, SCF MAX2 , and the transcriptional repressor protein SUPPRESSOR OF MAX2 1 (SMAX1), leading to the degradation of SMAX1 through the ubiquitin–proteasome system ( Liu et al ., 2014 ; Stanga et al ., 2013 ; Soundappan et al ., 2015 ; Yao et al ., 2017 ; Nelson, 2021 ). Strigolactone receptors, which belong to the α/μ-hydrolase superfamily, not only interact with SL, but also hydrolyze it ( Hamiaux et al ., 2012 ; Seto et al ., 2019 ). Recent biochemical and structural studies revealed that the SL hydrolysis intermediate covalently binds to AtD14 catalytic residues in the signaling complex, indicating that SL hydrolysis is indispensable for the activation of AtD14 ( Yao et al ., 2016 ; Bürger and Chory, 2020 ). However, the AtD14 D218A mutant, which has no SL hydrolysis activity because it lacks the catalytic triad, retained the ability to interact with AtSMXL7 in a SL-dependent manner and restored the atd14 mutant phenotype ( Seto et al ., 2019 ). Additionally, studies using chemical and biological methods indicated that the full activation of SL receptors depends on the hydrolysis of SL agonists, but the hydrolysis process is not necessarily required for SL signal transduction ( Wang et al ., 2022 ; Uraguchi et al ., 2018 ; Yasui et al ., 2019 ). Despite these advances, the exact function of SL hydrolysis and the timing of the formation of the complex are not completely understood. Notably, an analysis of the dynamics of SL-dependent complex formation associated with the hydrolysis process would provide new insights into the SL signaling mechanism. So far, the PPIs of SL signaling components have been detected using yeast-two hybrid, pull-down, and AlphaScreen assays. However, these methods are not suitable for kinetic analyses ( Zhao et al ., 2015 ; Wang et al ., 2021 ; Hamiaux et al ., 2012 ; Zhou et al ., 2013 ). Here, we developed the Time-Resolved Förster Resonance Energy Transfer (TR-FRET) assay system to quantitatively and kinetically detect the formation of the SL signaling complexes derived from A. thaliana and O. minor . The TR-FRET signal arises from the proximity between a luminescent lanthanide complex and a typical fluorescent dye (e.g. fluorescein, Cy5, or GFP). The physical properties of the lanthanide complexes, which have long lifetimes, sharp emission spectra, and large Stokes shift, enable the quantitative and kinetic analysis of PPIs, as well as the characterization of agonist/antagonist activities. Using the TR-FRET assay system, we were able to analyze the mode of action of SL receptor ligands. In addition, this method revealed that the SL receptor–repressor complex is so kinetically stable that an antagonist could not disrupt the PPIs within the already-formed complex. Time-course PPI analyses exhibited variations in intensity and time-dependency of the PPI signals among various SL analogs with different degrees of resistance to receptor-mediated hydrolysis. Overall, our results show that this novel technique can provide in-depth insights into the molecular mechanisms of the formation of the SL receptor complex. In addition to its use in mechanism-of-action studies, the TR-FRET assay system can rapidly and specifically detect naturally occurring SLs in root exudate mixtures. This selective activity-based probing method will facilitate the discovery of natural SL receptor ligands that control the signaling pathway. Results Development of the TR-FRET assay system to detect SL signaling complex formation To monitor the PPIs of SL signaling components, we set out to develop the TR-FRET assay system using the protein pairs AtD14–AtMAX2 and AtD14–AtSMXL7. We chose N- or C-terminal mEGFP-fused AtD14 with a 6×HN tag (mEGFP-AtD14 or AtD14-mEGFP, respectively) as the acceptor for the TR-FRET assay. The N-terminal 6×His-3×FLAG-tagged AtMAX2 was transiently expressed in Nicotiana benthamiana and purified by Co 2 + -affinity chromatography (Figure S1A,B). Based on a previous report, we expressed N-terminal 3×FLAG- and C-terminal 6×His-tagged AtSMXL7 in Escherichia coli , targeted to the periplasmic fraction using the pelB signal sequence (Figure S1C,D) ( Khosla et al ., 2020 ). Although the purified samples contained some impurities as well as AtMAX2/AtSMXL7, the pull-down assays showed that AtD14–AtMAX2 and AtD14–AtSMXL7 interactions proceeded in the presence of racemic GR24 ( rac -GR24, a mixture of (+)-GR24 and (−)-GR24), a synthetic agonist of the SL receptor ( Figure 1A ). These results confirm that the prepared proteins retained their original activities. Download figure Open in new tab Figure 1. (A) Pull-down assay to detect SL-induced complex formation of AtD14– AtMAX2 and AtD14–AtSMXL7 pairs. AtD14-mEGFP was detected by anti-GFP antibody. AtMAX2 and AtSMXL7 were detected by anti-FLAG antibody. IP, immunoprecipitation. WB, western blotting. (B) Schematic of TR-FRET assay for measuring SL-induced PPIs. The receptor protein is fused with mEGFP (GFP). The F-box and repressor proteins are FLAG-tagged and labeled by anti-FLAG antibody harnessing terbium-cryptate (Tb). SL induces each signaling complex resulting in a TR-FRET signal enhancing. (C) The dependency of each SL-signaling component in AtD14–AtMAX2 and AtD14–AtSMXL7 TR-FRET assays. Heatmap-projected values represent fold changes of TR-FRET signal upon treatment with rac -GR24 (50 µM) at a 300 min incubation period. Data are the means ± SD ( n = 3). Values of TR-FRET signals before normalization are shown in Figure S2A,B. Next, we performed TR-FRET assays using the same proteins at a range of concentrations. In these assays, we conducted in situ labeling of 3×FLAG-tagged AtMAX2 or AtSMXL7 with anti-FLAG antibodies connecting the terbium-cryptate FRET donors ( Figure 1B ). We measured TR-FRET signals, that indicate the ratio of the fluorescence intensity of the donor Tb-cryptate (485 nm) to that of the acceptor mEGFP (520 nm) in the absence or presence of rac -GR24 (Figure S2A,B). Figure 1C displayed TR-FRET signals upon addition of rac -GR24 normalized by the buffer control. The rac -GR24-dependent signals clearly increased in the presence of almost all pairs of proteins ( Figure 1C ). We found that AtD14-mEGFP yielded higher SL-dependent TR-FRET signals than mEGFP-AtD14 with both partner proteins ( Figure 1C ). In the presence of AtD14-mEGFP (100 nM), the SL-dependent TR-FRET signal slightly increased from 1.23 to 1.57 upon addition of AtMAX2 ( Figures 1C ). In the case of the AtD14–AtSMXL7 pair, the addition of AtSMXL7 raised the SL-dependent signal from 1.13 to 2.59 in the presence of 100 nM of AtD14-mEGFP ( Figure 1C ). In contrast, partner protein-independent nonspecific signals were observed in the presence of 500 nM AtD14. In the following experiments, therefore, we used AtD14-mEGFP (100 nM) as the acceptor because sufficient SL-triggered PPI signals were detected with this concentration. To confirm that the signal increase in the presence of rac -GR24 was derived from the structural activation of AtD14, TR-FRET assays were performed by using the catalytic triad mutant, AtD14 S97A , which has been reported to lose both the catalytic and signaling function ( Hamiaux et al ., 2012 ; Seto et al ., 2019 ). No signal induction was observed in either AtD14 S97A –AtMAX2 or the AtD14 S97A –AtSMXL7 upon addition of enough rac -GR24 (Figure S3A,B). To further examine the specificity of these signals, we prepared 6×His-AtMAX2 and StrepII-HiBiT-AtSMXL7-6×His without the anti-FLAG epitope tags (Figure S3C,D,E,F). As expected, the SL-dependent PPI signals were not increased in a protein-dependent manner, even in the presence of Tb cryptate-modified anti-FLAG antibodies (Figure S3G,I). In addition, these negative control proteins competed with the 3×FLAG-tagged AtMAX2 and AtSMXL7 (Figure S3H,J). These results show that our TR-FRET assay system specifically detects SL-dependent PPI behavior in AtD14–AtMAX2 and AtD14–AtSMXL7. We used the AtD14–AtSMXL7 pair in the following experiments because of its higher signal to noise ratio compared with that of AtD14–AtMAX2. Quantitative and kinetic analysis of complex formation by the TR-FRET assay system It has been reported that (+)-GR24, the 2 ’R stereoisomer of GR24, more strongly inhibits shoot branching in SL biosynthesis mutants of Arabidopsis than does (−)-GR24, the 2 ’S stereoisomer ( Scaffidi et al ., 2014 ). Despite the distinct biological activities of the enantiomer pair, in in vitro analyses, namely, differential scanning fluorometry (DSF), there were no significant differences in the thermal stability shift in the presence of each stereoisomer (Figure S4), consistent with the results of a previous study ( Scaffidi et al ., 2014 ; Waters et al ., 2015 ). To investigate the difference in the mode of action between (+)-GR24 and (−)-GR24, quantitative and kinetic TR-FRET assays using the AtD14–AtSMXL7 pair were carried out. Time-course measurements showed gradual increases in PPI signals in both cases ( Figure 2A ). However, the maximum intensity of the PPI signal was significantly different between (+)-GR24 and (−)-GR24, in spite of their comparable 50% effective concentration (EC50) values (0.30 and 0.68 µM, respectively) ( Figure 2B ). These results show that (−)-GR24 hardly activates the PPI, although it is recognized by the receptor. Download figure Open in new tab Figure 2. Quantitative and kinetic analysis of the AtD14 agonist and antagonist activities in TR-FRET assays. (A) Kinetic measurements of AtD14–SMXL7 TR-FRET signals in the presence of indicated concentration of (+)-GR24 and (−)-GR24. The assays were performed using AtD14-mEGFP (100 nM) and 3×FLAG-AtSMXL7 (5 µg/mL). Data are the means ± SE ( n = 3). (B) Dose-titration of (+)-GR24 and (−)-GR24 in AtD14–AtSMXL7 TR-FRET assays at the single time point (300 min) extracted from the data as shown in Figure 2A . Dots represent individual data values of three technical replicates. (C) Kinetic analysis of antagonistic activity of tolfenamic acid on GR24-induced AtD14–AtSMXL7 complex formation. The assay was performed using AtD14-mEGFP (100 nM) and 3×FLAG-AtSMXL7 (5 µg/mL). Control was performed without (+)-GR24 and tolfenamic acid. Data are the means ± SE ( n = 3). (D) Dose-response curve of tolfenamic acid in TR-FRET inhibition assay with GR24-induced AtD14–AtSMXL7 at the single time point (300 min) extracted from the data as shown in Figure 2C . Dots represent individual data values of three technical replicates. (E) The antagonistic effect of tolfenamic acid after the formation of AtD14–AtSMXL7 complex. The red arrow indicates the timing of tolfenamic acid treatment. The assay was performed using AtD14-mEGFP (100 nM) and 3×FLAG-AtSMXL7 (5 µg/mL). Data are the means ± SE ( n = 3). Next, to evaluate the applicability of the TR-FRET assay system, we investigated the antagonistic activity of tolfenamic acid, which is an antagonist of the SL receptors of petunia, rice, and Arabidopsis ( Hamiaux et al ., 2018 ). When tolfenamic acid was supplied alongside (+)-GR24 to the AtD14–AtSMXL7 pair, the increase in the TR-FRET signal was clearly inhibited ( Figure 2C ). As shown in Figure 2D , the 50% inhibitory concentration (IC50) value at 300 min was calculated as 1.65 µM, similar to the Ki value against AtD14 (2.52 µM) ( Hamiaux et al ., 2018 ). To further examine whether this SL receptor antagonist could disrupt the already-formed SL signaling complex, tolfenamic acid was added after the protein pair was pre-incubated with (+)-GR24 for 120 min. Surprisingly, while the increase in the TR-FRET signal stopped immediately upon addition of the antagonist, the elevated signal barely decreased ( Figure 2E ). These results suggest that the AtD14–AtSMXL7 complex is kinetically stable and/or the antagonist cannot access the catalytic pocket of the receptor in the ternary complex. Adaptation of TR-FRET assay system to O. minor SL signaling components Next, to expand the applicability of this system, we developed the TR-FRET assay system for the O. minor receptor–repressor protein pair. Another study conducted a functional analysis of SL receptors in O. minor , and found that OmKAI2d3, one of the KAI2d gene products, was highly sensitive to SL ( Takei et al ., 2023 ). We considered that quantitative and kinetic TR-FRET analyses would provide new insights into this sensitivity. Because the SMAX1 homolog in O. minor has not been identified, we searched for a SMAX1-like gene (KAL6540832.1) in the entire O. minor genome ( Bürger et al ., 2025 ). As shown in the phylogenetic tree, the amino acid sequence of this gene product had high homology with SMAX1 and SMXL2 from A. thaliana (Figure S5). Thus, we named this gene OmSMAX1 and cloned its coding sequence from the cDNA of O. minor . Initially, 6×HN-tagged OmKAI2d3-mEGFP and 3×FLAG-OmSMAX1-6×His were prepared in the same way as the corresponding proteins of A. thaliana, but almost all of the full-length OmSMAX1 protein was degraded after affinity column purification (Figure S6A,B). Because the previous study showed that the D1M-domain of SMAX1 is an important region for ligand-induced PPI, we prepared the D1M domain of OmSMAX1 (OmSMAX1-D1M) as an alternative to the full-length protein to improve its stability (Figure S6C,D) ( Khosla et al ., 2020 ). The results of a pull-down assay showed that the PPI between OmKAI2d3 and OmSMAX1-D1M was enhanced by rac -GR24, indicating that the D1M domain is also a critical region for the SL-dependent PPI between OmKAI2d3 and OmSMAX1 ( Figure 3A ). Next, we performed the TR-FRET assay using these proteins and found that SL-dependent TR-FRET signals significantly increased in OmKAI2d3- and OmSMAX1-D1M-dependent manners ( Figure 3B , Figure S7). In the presence of 100 nM OmKAI2d3-mEGFP and 5 µg/mL 3×FLAG-OmSMAX1-D1M, the TR-FRET signal increased 2.36-fold upon rac -GR24 treatment, with a low OmSMAX1-D1M-independent signal. Accordingly, we used these conditions for subsequent experiments. Download figure Open in new tab Figure 3. Adaptation of the TR-FRET assay system to the homologs’ pair of O. minor. (A) Pull-down assay to detect SL-induced complex formation between OmKAI2d3 and OmSMAX1 D1M domain. OmKAI2d3-mEGFP was detected by anti-GFP antibody. OmSMAX1-D1M was detected by anti-FLAG antibody. IP, immunoprecipitation. WB, western blotting. (B) The dependency of each SL-signaling component in TR-FRET assays. Heatmap-projected values represent fold changes of TR-FRET signal upon treatment with rac -GR24 (50 µM) at a 300 min incubation period. Data are the means ± SD ( n = 3). Values of TR-FRET signals before normalization are shown in Figure S7. The data in the absence of OmSMAX1 are identical to those in Figure 9S. To further validate the TR-FRET profiles, we performed the assays using (+)-GR24 and (−)-GR24. The results showed that (+)-GR24 more strongly increased the PPI signal of OmKAI2d3–OmSMAX1-D1M than did (−)-GR24 in a time-dependent manner ( Figure 4A ). As measured at a single time point, the EC50 value of (+)-GR24 for complex formation was 0.046 µM, and that of (−)-GR24 was about 1000-times higher ( Figure 4B ). Previous studies showed that the 2’ R configuration at the methyl butenolide ring is important for the agonistic activity of SL analogs on germination of root parasitic plants ( Umehara et al ., 2015 ; Scaffidi et al ., 2014 ). Consistent with the results of previous studies, our in vivo germination assays indicated that our O. minor seeds responded more strongly to (+)-GR24 than to (−)-GR24 (Figure S8A). To analyze the direct interaction between these chemicals and OmKAI2d3, we performed a DSF assay. The results showed that (+)-GR24 significantly decreased the melting temperature of OmKAI2d3, but (−)-GR24 did not (Figure S8B). These results demonstrate that our TR-FRET assay system can specifically and quantitatively evaluate the activity of SL agonists to induce PPIs among the SL signaling components of O. minor, just like in A. thaliana . Furthermore, the TR-FRET analysis revealed that OmKAI2d3 is activated by (+)-GR24 with affinity in the tens of nanomolar range, suggesting that the strong ligand perception ability of OmKAI2d3 is responsible for the SL hypersensitivity of O. minor seeds. Download figure Open in new tab Figure 4. Quantitative and kinetics analyses of the O. minor SL signaling complex formation triggered by various agonists. (A) Kinetic measurements of OmKAI2d3–OmSMAX1-D1M TR-FRET signals in the presence of indicated concentration of (+)-GR24 and (−)-GR24. The assay was performed using OmKAI2d3-mEGFP (100 nM) and 3×FLAG-OmSMAX1-D1M (5 µg/mL). Data are the means ± SE ( n = 3). (B) Dose-titration of (+)-GR24 and (−)-GR24 in OmKAI2d3–OmSMAX1-D1M TR-FRET assay at the single time point (300 min) extracted from the data as shown in Figure 4A . Dots represent individual data values of three technical replicates. (C) Kinetic analyses of agonistic activity of No.61 and 2,4-D-SL on OmKAI2d3–OmSMAX1-D1M complex formation. The assays were performed using OmKAI2d3-mEGFP (100 nM) and 3×FLAG-OmSMAX1-D1M (5 µg/mL). Data are the means ± SE ( n = 3). (D) Dose-titration of No.61 and 2,4-D-SL in OmKAI2d3–OmSMAX1-D1M TR-FRET assay at the single time point (300 min) extracted from the data as shown in Figure 4C . Dots represent individual data values of three technical replicates. (E) Kinetic comparison of TR-FRET signals upon addition of the SL receptor agonist (10 µM each) extracted from Figure 4A,C . Kinetic analysis of the OmKAI2d3–OmSMAX1 interaction using various SL analogs We found that the TR-FRET signal increased in time-dependent manner upon treatment with (+)-GR24 over the time scale of hours. Considering the hydrolytic activity of SL receptors, we hypothesized that the time-dependency of the SL-induced PPI signal is associated with the hydrolysis of SL agonists. To investigate the relationship between SL hydrolysis and PPI induction, we performed kinetic analyses of the PPI-inducing activities of three SL analogs, namely, (+)-GR24, No.61, and 2,4-D-SL. It has been reported that No.61 is more resistant to OmKAI2d3-mediated hydrolysis than is rac -GR24, and that No.61 strongly induces germination of O. minor ( Kawada et al ., 2024 ). In addition, it has been reported that 2,4-D-SL is more quickly hydrolyzed than is rac -GR24, and that 2,4-D-SL potently inhibits the hydrolysis of Yoshimulactone Green, a pro-fluorescent SL agonist ( Tsuzuki et al ., 2024 ). Despite its ability to bind to OmKAI2d3, 2,4-D-SL exhibited weaker activity to induce O. minor germination compared with rac -GR24. The TR-FRET assay showed that both No.61 and 2,4-D-SL induced the PPI of OmKAI2d3–OmSMAX1-D1M ( Figure 4C ). The EC50 value of No.61 was estimated to be 0.135 µM, consistent with its strong ability to induce O. minor germination ( Figure 4D ). In contrast, 2,4-D-SL did not elicit a sigmoidal dose-response curve ( Figure 4D ). Remarkably, we also found that maximum signal intensities differed depending on the type of SL agonist. As shown in Figure 4E , when comparing the time-dependent PPI signals among three agonists at 10 µM, the signal induced by 2,4-D-SL stopped earlier than did the signals induced by (+)-GR24 and No.61. Consequently, the maximum signal intensity induced by 2,4-D-SL was lower than those induced by (+)-GR24 and No.61 ( Figure 4E ). Considering its high susceptibility to OmKAI2d3-mediated hydrolysis, these results suggest that the rapid hydrolysis of 2,4-D-SL may prevent effective and stable induction of the PPI. Therefore, our TR-FRET assay system offers novel insights into PPI dynamics based on their momentary affinity, hydrolyzability, and receptor activation potential. Selective sensing of the naturally occurring SL receptor agonists from root exudate As described above, the TR-FRET assay system with the hypersensitive SL receptor OmKAI2d3 easily and quantitatively detected several hundred picograms of active SL in a 20-µL reaction volume. In general, the time-gated and ratiometric measurements in TR-FRET assays suppress interference from irrelevant compounds ( Bazin et al ., 2002 ). Thus, we speculated that this method would be a useful tool to detect naturally occurring SLs from crude samples containing diverse natural products. Although the full-length OmSMAX1 protein was degraded as mentioned above, we detected the SL-dependent TR-FRET signal from a mixture containing the full-length OmSMAX1 and many degradation products with a slightly higher signal to noise ratio that that of a mixture containing OmSMAX1-D1M (Figure S9). Accordingly, we used 3×FLAG-OmSMAX1-6×His in SL-sensing experiments. The FRET efficiency is influenced by the donor/acceptor wavelength, orientation, and distance. Terbium-cryptate is one of the most popular lanthanide complexes for the TR-FRET assays by combining with the various types of green- and red-fluorescent dyes as the acceptors. Thus, toward further optimization of the signal to nose ratio, we adapted the SNAP tag labeled by fluorescein (FL) or sulfo-cyanine5 (Sulfo-Cy5) instead of mEGFP ( Keppler et al ., 2003 ). Furthermore, as the structural basis of the complex between the SL receptor and the transcriptional repressor had never been experimentally revealed, we roughly tested two different linker lengths between OmKAI2d3 and mEGFP. Even though these were not significantly optimized conditions, we demonstrated the capability of the FL- or Sulfo-Cy5-labeled SNAP tag in addition to mEGFP (Figure S10A,B). In the following SL sensing experiments, we used the pair 3×FLAG-OmSMAX1-6×His and 6×HN-OmKAI2d3-GSSG-mEGFP, in which mEGFP was connected at the C-terminus by a Gly-Ser-Ser-Gly short linker. We performed TR-FRET assays using extracts of rice (cv. Shiokari) hydroponic cultures that contained numerous organic compounds, including SLs such as 4-deoxyorobanchol ( Figure 5A ) ( Umehara et al ., 2008 ). We found that root exudates of d14 , the rice mutant that strongly accumulates SL because it lacks the SL receptor, strongly increased the TR-FRET signal ( Figure 5B ). In contrast, root exudates of d10 , the rice SL biosynthesis mutant defective in endogenous SLs, barely generated a response ( Figure 5B ) ( Arite et al ., 2009 ; Umehara et al ., 2008 ; Arite et al ., 2007 ). Additionally, the addition of rac -GR24 to d10 root exudate restored the TR-FRET signal to the same level as that obtained with d14 root exudate, indicating that agonistic activities can be specifically detected even in crude samples ( Figure 5C ). In addition, the EC50 value of (+)-GR24 was hardly influenced by the d10 root exudate (Figure S11). These results clearly demonstrate the utility of this method for specifically detecting naturally occurring SLs with minimal interference. Download figure Open in new tab Figure 5. Application of the TR-FRET assay system for sensing the naturally occurring SLs. (A) Schematic of the high-throughput TR-FRET-based SL sensing system. Natural compound mixtures are extracted from root exudates and subjected to a 96-well TR-FRET assay format using the highly-sensitive OmKAI2d3–OmSMAX1 pair without further purification. (B) SL detection from root exudate of rice mutants ( d10 : SL deficient mutant, d14 : SL accumulated mutant) using the TR-FRET system. Each data represents the individual assay values of three biological replicates. (C) Complementation of weak TR-FRET signals in d10 root exudate by supplementing rac -GR24. The assays were performed using OmKAI2d3-GSSG-mEGFP (100 nM) and 3×FLAG-OmSMAX1 (5 µg/mL). Concentrations of root exudate samples are expressed as a fold enrichment relative to the original volume of the hydroponic culture media. Data are the means ± SE ( n = 3). Discussion We developed an assay system to monitor the formation of SL signaling complexes derived from A. thaliana and O. minor using TR-FRET technology. Additionally, we demonstrated that our TR-FRET assay system can distinguish the mode of action of different types of ligands for SL receptors. For example, assays with (+)-GR24 and (−)-GR24 exhibited remarkable differences in the maximum signal intensity, rather than EC50 values, of AtD14–AtSMXL7 complex formation. Considering the weak but significant activity of (−)-GR24 to inhibit branching in Arabidopsis, our results suggest that (−)-GR24 might function as a partial agonist for AtD14 ( Scaffidi et al ., 2014 ). It is noteworthy that our TR-FRET assay system defines the degree of agonistic activity semi-quantitatively, reflecting in vivo activity. Even though the mechanism of partial activation of AtD14 by (−)-GR24 is unclear, the weak induction of the TR-FRET signal suggests that the free energy landscape of AtD14 activation depends on the ligand configuration ( Chen and Shukla, 2023 ; Chen et al ., 2022 ). Our kinetic analyses revealed that, although adding an antagonist abolished the activation of AtD14, it did not disrupt the already-formed AtD14–AtSMXL7 complex. One possibility is that the dissociation of AtD14–AtSMXL7 is quite slow because of multi-point interactions on the protein–protein contact surface. Meanwhile, mass spectrometric and crystallographic analyses of SL-supplied receptors have demonstrated that the methyl butanolide ring is covalently trapped at the catalytic amino acid residue. Therefore, it is also possible that this covalent attachment prevents the antagonist from binding. In either case, the results provide new insights into the dynamics of the SL signaling complex, as well as the importance of the timing of antagonist treatment. The kinetic TR-FRET assay showed the signal induction by 2,4-D-SL decreased faster than that by (+)-GR24 in the OmKAI2d3–OmSMAX1-D1M complex. This observation indicates that excessively hydrolysable SL analogs cannot sufficiently activate SL receptors for complex formation, possibly because the receptor-mediated hydrolytic deactivation of these chemicals is too fast. Accordingly, although we have not yet concluded whether the PPI occurs before or during the hydrolysis process, strong tolerance of SL agonists to receptor-mediated hydrolysis would contribute to the effective activation of the SL signal. We observed slight increases in TR-FRET signals even in the absence of agonists in the AtD14–AtSMXL7 and OmKAI2d3–OmSMAX1-D1M pairs, but not in the AtD14–MAX2 pair (Figure S2B; Figure S7). The trend was also observed in our pull-down assays ( Figure 1A ; Figure 3A ) as well as in previous studies ( Yao et al ., 2017 ; Jiang et al ., 2013 ). These results indicate that the receptor proteins can partially recruit the repressor proteins independently of SL. To prevent inappropriate activation of SL signaling via degradation of repressors, the recruitment of the E3 ubiquitin ligase that includes MAX2 is likely strictly controlled by SL. Furthermore, we should investigate whether the SL-independent receptor–repressor interaction occurs in planta and influences the regulation of SL signaling. This is especially important for obligate parasitic plants because tight control of the signal repression mechanism is crucial to prevent death via undesired germination in the absence of host plants. The dynamics of PPIs in receptor complexes of the plant hormone auxin have been analyzed using surface plasmon resonance (SPR), providing critical insights into the molecular mechanism of auxin signaling (Calderón Villalobos et al ., 2012 ). For example, different auxins differentially induce the formation of auxin co-receptor complexes, as measured by their association and dissociation rates (Calderón Villalobos et al ., 2012 ). In addition, the binding kinetics differ among the auxin co-receptor pairs, which are highly duplicated in angiosperms (Calderón Villalobos et al ., 2012 ). As shown in studies on auxin, information on the kinetic and thermodynamic characteristics of ligand-induced PPIs is crucial not only for understanding the biological functions of plant hormones, but also for the development of highly potent ligands. However, few studies have explored the kinetics of ligand-induced PPIs under the research of other plant hormones. TR-FRET is a promising high-throughput platform to quantitatively and kinetically characterize molecular interactions, and it is widely used in the field of drug discovery (Greenway et al ., 2024; Falkenstern et al ., 2024). In this study, we developed a kinetic assay system for monitoring the formation of the SL signaling complex using TR-FRET, offering a novel strategy for future research in SL signaling. In addition, our study provides a technical basis for establishing a similar kinetic assay format for other plant hormones that induce PPIs ( Takeuchi et al ., 2021 ). We also demonstrated that our TR-FRET assay can selectively detect naturally occurring SLs from plant-derived samples containing numerous natural products. Compared with conventional liquid chromatography–tandem mass spectrometry and phenotype-based SL detection, the TR-FRET system has advantages of rapid and activity-based detection of SL molecules, as well as compatibility with impurities that inhibit plant growth and development. We also demonstrated the use of the SNAP tag, which is easily labeled by FL and Sulfo-Cy5, as the acceptor in our TR-FRET assay system. The multi-color platform will enable robust screening with validation of true fractions containing active compounds ( Hassig et al ., 2014 ). The structures of SLs are diversified among plant species, and more than 30 SL derivatives have been discovered ( Chesterfield et al ., 2020 ). However, the active hormone in the shoot-branching pathway has not been identified ( Seto, 2023 ). We are currently working to detect and isolate unknown active hormones related to the inhibition of shoot branching, as well as diverse natural compounds with SL-like activities that are biosynthesized in various species. Surprisingly, we observed a fast and strong increase in the PPI signal upon addition of d14 root exudate, compared with that obtained using synthetic rac -GR24 as an authentic control. Moreover, the addition of rac -GR24 to d10 root exudate restored the PPI signal to the same level as that obtained with d14 root exudate. Rather than differences in activity between GR24 and natural SLs, these results suggest that rice root exudate contains putative enhancers, such as cofactors or allosteric modulators, for the formation of the OmKAI2d3–OmSMAX1 complex. Previous crystallographic and biochemical studies indicate that auxin and jasmonate receptors require inositol polyphosphates as essential cofactors ( Sheard et al ., 2010 ; Tan et al ., 2007 ). Likewise, in the SL signaling complex, the structural dynamics of MAX2 are controlled by citrate, a widely distributed organic acid, and the citrate-dependent conformational switch of MAX2 is thought to be associated with SL signal transduction ( Tal et al ., 2022 ; Shabek et al ., 2018 ). Thus, it is possible that there are modulators of the SL receptor–repressor interaction, although further research is needed to identify and functionally characterize them. Application of the TR-FRET system for other plant hormones will be useful to identify unknown chemicals that control plant hormone signaling. Notably, despite previous attempts to identify the endogenous ligand for KAI2, the chemical identity of this plant hormone candidate has not yet been discovered (Waters and Nelson, 2022). Given the high structural similarity of KAI2 to SL receptors such as OmKAI2d3 and AtD14, it will be feasible to develop a KAI2-based TR-FRET system that can be used to discover unknown KAI2 ligands. In summary, we developed a novel method based on TR-FRET to monitor the formation of SL signaling complexes. This kinetic and high-throughput assay system will facilitate further research on the dynamics of important molecular interactions among SL signaling components. In addition, activity-based screening using this method will allow for the discovery of potent SL signaling modulators from biological resources and drug-like chemical libraries. Materials and Methods Chemicals and general methods Tolfenamic acid was obtained from Tokyo Chemical Industry Co., Ltd. (Tokyo, Japan). (+)-GR24 and (−)-GR24 were prepared as described in supporting information. No.61, 2,4-D-SL, and the fluorescein O 6 -benzylguanine conjugate (FL-BG) were synthesized as described in previous papers ( Kawada et al ., 2024 ; Tsuzuki et al ., 2024 ; Ramil et al ., 2017 ). The sulfo-Cy5 O 6 -benzylguanine conjugate (Sulfo-Cy5-BG) was synthesized as described in supporting information. All reagents for chemical syntheses were purchased from Tokyo Chemical Industry Co., Ltd. (Tokyo, Japan) and FUJIFILM Wako Pure Chemical Corp. (Osaka, Japan). NMR spectra were measured by a NMR spectrometer JNM-ECZL (JEOL). High-resolution mass spectrometry (HRMS) was performed by a quadrupole/time-of-flight tandem mass spectrometer X500R (AB SCIEX). Vector construction To substitute the 6×His tag of pET47b to 6×HN tag, the hybridized DNA oligos coding 6×HN tag, described in Table S1, was cloned into linearized pET47b at the site between NdeI and SacⅡ using the In-Fusion cloning Kit (Takara). Hereafter, we referred to this vector as pET47b-6×HN. To prepare vectors fusing mEGFP and SNAP tag at C-terminus, each artificial DNA fragment (Eurofins Genomics) was inserted into the site between XhoI and PacI of pET47b-6×HN based on In-Fusion cloning. For the N-terminal tagging of mEGFP and SNAP, each coding sequence was amplified by PCR and individually inserted into the site between SmaI and SacI. For preparing the protein expression vectors of AtD14 and OmKAI2d3 connecting mEGFP/SNAP based on their modified pET47b-6×HN, the coding sequences of AtD14, AtD14 S97A and OmKAI2d3, which have been used in our previous studies ( Seto et al ., 2019 ; Takei et al ., 2023 ), were amplified by PCR with primers described in Table S1. Each PCR product was introduced into the site between SmaI–SacI and XhoI–PacI for the N-terminal and C-terminal tagging, respectively. To substitute the original linker (the sequence of thrombin and S-tag) in the middle of OmKAI2d3 and mEGFP with Gly-Ser-Ser-Gly (GSSG) linker, the annealed DNA fragment as shown in Table 1 was cloned into the linearized pET47b-6×HN vector carrying OmKAI2d3 and mEGFP digested at SacI and XhoI sites. The protein expression vectors for AtSMXL7 and OmSMAX1(-D1M) were constructed by using pET26b, which encodes pelB signal sequence and C-terminal 6×His tag, according to the previous study ( Khosla et al ., 2020 ), with slight modifications. The hybridized 3×FLAG tag-coding oligos described in Table S1 was cloned into the site between NcoI and BamHI of pET26b by In-Fusion cloning, yielding pET26b-3×FLAG. The cording sequence of AtSMXL7 ( Seto et al ., 2019 ) was amplified by PCR with primers described in Table S1. The cording sequence of OmSMAX1 which is predicted from genome data of O. minor ( Bürger et al ., 2025 ) was amplified from cDNA of O. minor by PCR with primers described in Table S1. The region of OmSMAX1-D1M (A164 to K610) was determined according to the previous study ( Khosla et al ., 2020 ). Each DNA fragment was cloned into the site between BamHI and XhoI of pET26b-3×FLAG by In-Fusion cloning. For the construction of StrepⅡ-HiBiT-AtSMXL7-6×His protein expression vector, the insert DNA fragment of StrepⅡ-HiBiT was prepared by primer extension using partially complemental oligos described in Table S1. The elongated DNA fragment was cloned into pET26b by In-Fusion cloning. The following operations were carried out as described above. The CDSs of AtMAX2 and ASK1 ( Seto et al ., 2019 ) were amplified by PCR with primers described in Table S1. For the transient co-expression of 6×His-AtMAX2 and tag-free ASK1 in tobacco leaves, these PCR products were sub-cloned into pDONR207 by BP reaction and recombined into pEAQ-HT DEST 2 and pEAQ-HT DEST1 for AtMAX2 and ASK1, respectively ( Sainsbury et al ., 2009 ). For 6×His-3×FLAG-AtMAX2 expression, the plasmid was prepared following GoldenBraid assembly systems previously developed (Sarrion-Perdigones et al ., 2013). The GoldenBraid 2.0 kit was a gift from Diego Orzaez (Addgene kit # 1000000076). The entry vectors of 6×His-3×FLAG for insertion at B3 site and of AtMAX2 for that at B4–B5 site were sub-cloned into pUPD2 vector via Golden Gate cloning by using BsmBI restriction enzyme and DNA ligase (New England Biolabs). Then, sequences of Arabidopsis Ubiquitin10 promoter (GoldenBraid 2.0 kit), 6×His-3×FLAG, AtMAX2, and Arabidopsis Ubiquitin3 terminator (GoldenBraid 2.0 kit) within pUPD2 were assembled into pDGB3 vector by BsaI restriction enzyme and DNA ligase (New England Biolabs). Protein expression and purification AtD14, AtD14 S97A and OmKAI2d3 protein connecting mEGFP or SNAP were purified as previously described and stored at –80°C until use ( Tsuzuki et al ., 2024 ). The expression and purification of AtSMXL7 and OmSMAX1 proteins was performed according to the previous study ( Khosla et al ., 2020 ) with slight modifications. After protein expression induction, the E. coli cells were harvested by centrifugation at 3,700 × g and the pellet were stored at −30℃ until use. The pellet was resuspended in lysis buffer [50 mM sodium phosphate buffer (pH 7.6), 300 mM NaCl, 5 mM imidazole, 1× EDTA-free protease inhibitor cocktail (nacali tesque), 1.5 mg/mL Lysozyme, 0.1% Tween 20, with or without 10 unit/mL Benzonase (Merck)] and incubated for 30–60 min on ice. The resuspended cells were lysed with gentle sonication. After centrifugation at 18,000 × g for 30 min, the supernatant was subjected affinity chromatography using nickel sepharose resin. The column was washed with the washing buffer [50 mM sodium phosphate buffer (pH 7.6), 300 mM NaCl, 5–20 mM imidazole]. The bound protein was eluted with the elution buffer [50 mM sodium phosphate buffer (pH 7.6), 300 mM NaCl, 250 mM imidazole]. The eluate was concentrated using VIVASPIN Turvo15 (Sartorius) and buffer-exchanged into 1× PBS (–). The protein solution was aliquoted to the appropriate volume, immediately frozen in liquid nitrogen, and stored at −80℃ until use. OmSMAX1-D1M was prepared by the same procedure as described above with slight changes in buffers. Lysis buffer [50 mM sodium phosphate buffer (pH 7.0), 500 mM NaCl, 1× EDTA-free protease inhibitor cocktail, 1.5 mg/mL Lysozyme, 0.1% Tween 20, 10 unit/mL Benzonase], washing buffer [50 mM sodium phosphate buffer (pH 7.0), 300 mM NaCl, 20 mM imidazole], and elution buffer [50 mM sodium phosphate buffer (pH 7.0), 300 mM NaCl, 200 mM imidazole] were employed for OmSMAX1-D1M purification. For the expression of AtMAX2 protein, the expression vectors containing AtMAX2 gene and ASK1 were introduced into Agrobacterium tumefacience LBA4404, respectively. The transformed Agrobacterium cells were pre-cultured in LB selection medium at 30℃ with shaking. Overnight cultures were diluted into fresh LB medium and incubated at 30℃ until the OD600 reached to 0.5–1.0. The Agrobacterium cells were harvested by centrifugation at 3,500 × g and re-suspended in infiltration buffer [10 mM MES-KOH (pH 5.6), 150 mM acetosyringone, 10 mM MgCl2] to adjust OD600 to 0.5-1.0. After placing under dark conditions for at least 1 h, the Agrobacterium solutions for AtMAX2 and ASK1 expression were mixed (1:1) and infiltrated into N. benthamiana leaves. The leaves were harvested at 5-7 days post-infiltration. 10 g of the leaves were ground in a mortar with the aid of liquid nitrogen. The powdered leaves were extracted with 15 mL of the extraction buffer [50 mM HEPES-NaOH (pH 8.0), 150 mM NaCl, 5 mM 2-mercaptoethanol, 50 µM MG-132, 10% glycerol, 0.5% Triton X-100, 1× EDTA-free protease inhibitor cocktail]. After the centrifugation at 18,000 × g for 15 min, the supernatants were filtered through Miracloth (Merck). The filtrate was centrifuged at 18,000 × g for 15 min. The supernatant was subjected affinity chromatography using TALON superflow resin (Cytiva). The column was washed with the washing buffer [50 mM HEPES-NaOH (pH 8.0), 150 mM NaCl, 5 mM imidazole, 0.5% Triton X-100). The bound protein was eluted with the 1×PBS (–) containing 150 mM imidazole. The eluate was concentrated using VIVASPIN Turvo15 (Sartorius) and buffer-exchanged into 1× PBS (–). The protein solution was aliquoted to the appropriate volume, immediately frozen in liquid nitrogen, and stored at −80℃ until use. Fluorescent labelling of SNAP tag SNAP-tagged proteins (50 µM) were incubated with FL-BG (200 µM) or Sulfo-Cy5-BG (200 µM) in 1× PBS (–) containing 1mM DTT at 25℃ for 2 h. Un-reacted labelling regents were removed by using desalting column (PD-10; Cytiva). After concentrating protein solutions by VIVASPIN, the concentration of each protein was determined by the Bradford method. The labeling yield were calculated from the absorbance (FL: the molar extinction coefficient of 68,000 M −1 cm −1 at 494 nm; Sulfo-Cy5: the molar extinction coefficient of 250,000 M −1 cm −1 at 649 nm) and estimated as >70%. In vitro Pull-down assay AtD14-mEGFP (25 µg) was immobilized on 10 µL of GFP-Trap Magnetic Particles M-270 (Proteintech) at 4℃ for 1 h with agitation. The beads were washed with 500 µL of washing buffer [50 mM MES-NaOH (pH6.5), 150 mM NaCl, 0.1 mg/mL BSA, 10% glycerol, 0.1 % Tween 20] for 3 times. AtSMXL7 protein (45 µg) or AtMAX2 protein (20 µg) was added to the beads in 40 µL of assay buffer [50 mM MES-NaOH (pH6.5), 150 mM NaCl, 2 mM DTT, 0.1 mg/mL BSA, 10% glycerol, 1% DMSO, 0.1 % Tween 20] with or without 50 µM GR24. After incubation at room temperature for 1 h with agitation, the beads were washed by 200 µL of washing buffer for 5 times. Bound proteins were extracted by 20 µL of 1×SDS-sample buffer with boiling. The samples were subjected to SDS-PAGE and western blotting analysis. OmKAI2d3-mEGFP (12.6 µg) was immobilized on 10 µL of GFP-Trap Magnetic Particles M-270 at 4℃ for 1 h with agitation. The beads were washed with 500 µL of washing buffer [50 mM MES-NaOH (pH6.5), 150 mM NaCl, 0.1 mg/mL BSA, 10% glycerol, 0.1 % Tween 20] for 3 times. OmSMAX1-D1M protein (0.75 µg) was added to the beads in 40 µL of assay buffer [50 mM MES-NaOH (pH6.5), 150 mM NaCl, 2 mM DTT, 0.1 mg/mL BSA, 10% glycerol, 1% DMSO, 0.1% Tween 20] with or without 50 µM rac -GR24. After incubation at 4℃ for 30 min with agitation, the beads were washed by 200 µL of washing buffer for 5 times. Bound proteins were extracted by 20 µL of 1×SDS-PAGE sample buffer with boiling. The samples were subjected to SDS-PAGE and western blotting analysis. Western blotting Protein samples were separated by SDS-PAGE and transferred to a nitrocellulose membrane by semi-dry blotting system (ATTO). For the FLAG-tagged, His-tagged and mEGFP-tagged proteins, the membrane was blocked in 1×TBS-T containing 0.5% BSA at room temperature for 1 h. These proteins were detected by using 1:2000 dilution anti-DDDDK (FLAG)-tag monoclonal antibody-HRP conjugate (Medical & Biological Laboratories), 1:5000 anti-6×His monoclonal antibody-HRP conjugate (FUJIFILM Wako Pure Chemical) and 1:2000 dilution anti-GFP polyclonal antibody-Alexa Fluor 488 conjugate (Invitrogen). Chemiluminescent detections were carried out using EzWestLumi one (ATTO) and images were obtained using the iBright CL 1500 (Invitrogen). The HiBiT-tagged AtSMXL7 was detected by using Nano-Glo HiBiT Blotting System (Promega) according to manufactures protocol. TR-FRET assay The experiments were performed in HTRF 96-well low-volume white plate (Revvity) in 20 µL of reaction buffer [50 mM MES-NaOH (pH6.5), 150 mM NaCl, 2 mM DTT, 0.1 mg/mL BSA, 1× HTRF mAb anti-FLAG Tb-Conjugate (Revvity)] with the indicated concentrations of proteins, chemicals and solvents (acetone or DMSO). Unless otherwise stated, the final concentration of acetone was adjusted to 1% (v/v). After the FLAG-tagged protein was incubated in reaction buffer at room temperature for 30 min, the SL receptor proteins and chemical solutions were added to a reaction mixture on ice. Measurements were taken on a microplate reader Infinite 200 PRO F Plex (TECAN) with the following settings (mEGFP and SNAP-FL: excitation 340(35)nm, emission A 520(10)nm, emission B 485(20)nm, 120 µs delay time, 200 µs integration time, 5 min interval; SNAP-Sulfo-Cy5: excitation 340(35)nm, emission A 665(8)nm, emission B 620(10)nm, 100 µs delay time, 200 µs integration time, 5 min interval). The TR-FRET signal was taken as the emission A/emission B intensity ratio. In the kinetic monitoring of the antagonist activity on the post-formed complex, 1 µL of tolfenamic acid solutions containing 5 % acetone was added to 20 µL of reaction solutions at 120 min post-PPI induction by (+)-GR24 (0.5 µM). Measurements were taken as described above. TR-FRET assay using rice root exudates The strigolactone receptor mutant ( d14-1 ) and biosynthesis mutant ( d10-1 ) of Oryza sativa (rice) were used for the SL sensing experiments ( Ishikawa et al ., 2005 ). After growing for 24 days in a hydroponic culture system under phosphate deficient conditions as previously described ( Umehara et al ., 2015 ), root exudates of rice plants were extracted with ethyl acetate. The organic phase was dried over Na2SO4 and concentrated under a stream of nitrogen. The extracts were dissolved in DMSO and subjected to a TR-FRET assay. The experiments were performed in the reaction buffer containing 5% DMSO using the above procedure. When supplying the rac -GR24 or (+)-GR24 to d10 root exudate, 0.5% or 1% acetone was added to all samples, respectively. Germination assay O. minor germination assays were performed as previously described ( Tsuzuki et al ., 2024 ). DSF assay AtD14 and OmKAI2d3 were cloned into pET47b-(+) vector and these proteins were prepared as previously described ( Tsuzuki et al ., 2024 ). AtD14 protein (5.5 µg) or OmKAI2d3 protein (10 µg) was dissolved in 20 µL of 1× PBS (–) buffer containing SYPRO Tangerine or SYPRO Orange (SigmaAldrich) and each SL analog with 1% (v/v) acetone on a 96-well PCR white plate (Roche). SYPRO Tangerine (×5000) was diluted to ×1.25 final concentration in a reaction mixture for AtD14. SYPRO Orange (×5,000) was diluted to ×0.625 final concentration for OmKAI2d3. The mixtures were heated from 20℃ to 95℃ with scanning the fluorescence intensity (Ex/Em: 483/640 nm for AtD14; Ex/Em: 483/610 nm for OmKAI2d3) using LightCycler480 (Roche). The first derivative curve was calculated from a plot of the fluorescence intensity against the temperature using the LightCycler480 Software. Data analysis and graphics For the calculations of EC50 and IC50 values, the plots were fitted by a four-parameter logistic model using the R drc package ( Ritz et al ., 2015 ). Graphs were drawn using the R ggplot2 package ( Wickham, 2016 ). Author contribution T.S. performed majority of experiments in this paper with guidance from K.N. and Y.S.; Y.K., C.S. and T.I. performed part of protein preparation and TR-FRET analyses; M.B. supported cloning of O . minor gene; J.R. and M.K. prepared part of plasmids with guidance from S.H; K.F and T.A. prepared No.61; K.N. synthesized FL-BG and Sulfo-Cy5-BG; K.N. and Y.S. designed research. All authors wrote the manuscript. Funding JST ACT-X (JPMJAX22BH to K.N.), JSPS KAKENHI (19K05852, 22H02276, 23H05409 and 24H00878 to Y.S.; 22K14788 to J.R.), JST FOREST Program (JPMJFR211S to Y.S.), Mitsubishi Foundation (to Y.S.) and Kato Memorial Bioscience Foundation (to Y.S.). Conflict of Interest Statement The authors have no conflicts of interest to declare. Short legends for Supporting Information Figure S1. Preparation of FLAG-tagged AtMAX2 and AtSMXL7 recombinant proteins. Figure S2. TR-FRET signals in AtD14‒AtMAX2 and AtD14‒AtSMXL7 pairs in the absence or presence of rac -GR24 before normalization. Figure S3. Negative control of TR-FRET assays using the catalytic mutant protein of AtD14 and FLAG-tag-free partner proteins. Figure S4. Melting peaks of 6×His-AtD14 in the absence and presence of (+)-GR24 or (−)-GR24. Figure S5. Phylogenetic tree of SMAX family. Figure S6. Preparation of FLAG-tagged OmSMAX1 and OmSMAX1-D1M recombinant proteins. Figure S7. TR-FRET signals in OmKAI2d3‒OmSMAX1-D1M pairs in the absence and presence of rac -GR24 before normalization. Figure S8. Effects of (+)-GR24 and (−)-GR24 on O. minor seed germination and the melting temperature of OmKAI2d3. Figure S9. TR-FRET assay using OmKAI2d3 and full-length OmSMAX1. Figure S10. Effects of fluorophore and linker variations on the SL-dependent TR-FRET signals. Figure S11. Effects of d10 root exudates on the EC50 value of (+)-GR24. Preparation of (+)-GR24 and (−)-GR24. Chemical synthesis of Sulfo-Cy5-BG. Table S1. Primers used in this study. Acknowledgments We thank Nagasawa Water Purification Plant for the corporation regarding O . minor sampling. We thank Dr. Jennifer Smith from Edanz ( https://jp.edanz.com/ac ) for editing a draft of this manuscript. Funder Information Declared JST ACT-X , JPMJAX22BH JSPS KAKENHI , 19K05852 , 22H02276 , 23H05409 , 24H00878 , 22K14788 JST FOREST Program , JPMJFR211S Mitsubishi Foundation, https://ror.org/02y2f3e22 Kato Memorial Bioscience Foundation, https://ror.org/054n63512 Refere n ces 1. ↵ Akiyama , K. , Matsuzaki , K.-I. and Hayashi , H . ( 2005 ) Plant sesquiterpenes induce hyphal branching in arbuscular mycorrhizal fungi . Nature , 435 , 824 – 827 . OpenUrl CrossRef PubMed Web of Science 2. ↵ Arite , T. , Iwata , H. , Ohshima , K. , Maekawa , M. , Nakajima , M. , Kojima , M. , Sakakibara , H. and Kyozuka , J . ( 2007 ) DWARF10, an RMS1/MAX4/DAD1 ortholog, controls lateral bud outgrowth in rice . Plant J ., 51 , 1019 – 1029 . OpenUrl CrossRef PubMed Web of Science 3. ↵ Arite , T. , Umehara , M. , Ishikawa , S. , Hanada , A. , Maekawa , M. , Yamaguchi , S. and Kyozuka , J . ( 2009 ) d14, a strigolactone-insensitive mutant of rice, shows an accelerated outgrowth of tillers . Plant Cell Physiol ., 50 , 1416 – 1424 . OpenUrl CrossRef PubMed Web of Science 4. ↵ Bazin , H. , Trinquet , E. and Mathis , G . ( 2002 ) Time resolved amplification of cryptate emission: a versatile technology to trace biomolecular interactions . J. Biotechnol ., 82 , 233 – 250 . OpenUrl CrossRef PubMed 5. ↵ Bürger , M. and Chory , J . ( 2020 ) The many models of strigolactone signaling . Trends Plant Sci ., 25 , 395 – 405 . OpenUrl CrossRef PubMed 6. ↵ Bürger , M. , Soco , C.F. , García , M.A. , Seto , Y. and Leyva , A. ( 2025 ) Three Orobanche genomes reveal distinct molecular repertoires in species with different host ranges . Plant Cell Physiol ., https://academic.oup.com/pcp/advance-article/doi/10.1093/pcp/pcaf047/8134335 7. ↵ Calderón Villalobos , L.I.A. , Lee , S. , De Oliveira , C., et al. ( 2012 ) A combinatorial TIR1/AFB-Aux/IAA co-receptor system for differential sensing of auxin . Nat. Chem. Biol ., 8 , 477 – 485 . OpenUrl CrossRef PubMed 8. ↵ Chen , J. , Nelson , D.C. and Shukla , D . ( 2022 ) Activation Mechanism of Strigolactone Receptors and Its Impact on Ligand Selectivity between Host and Parasitic Plants . J. Chem. Inf. Model ., 62 , 1712 – 1722 . OpenUrl PubMed 9. ↵ Chen , J. and Shukla , D . ( 2023 ) Effect of histidine covalent modification on strigolactone receptor activation and selectivity . Biophys. J ., 122 , 1219 – 1228 . OpenUrl PubMed 10. ↵ Chesterfield , R.J. , Vickers , C.E. and Beveridge , C.A . ( 2020 ) Translation of strigolactones from plant hormone to agriculture: Achievements, future perspectives, and challenges . Trends Plant Sci ., 25 , 1087 – 1106 . OpenUrl CrossRef PubMed 11. ↵ Conn , C.E. , Bythell-Douglas , R. , Neumann , D. , et al. ( 2015 ) PLANT EVOLUTION. Convergent evolution of strigolactone perception enabled host detection in parasitic plants . Science , 349 , 540 – 543 . OpenUrl Abstract / FREE Full Text 12. ↵ Cook , C.E. , Whichard , L.P. , Turner , B. , Wall , M.E. and Egley , G.H . ( 1966 ) Germination of Witchweed (Striga lutea Lour.): Isolation and Properties of a Potent Stimulant . Science , 154 , 1189 – 1190 . OpenUrl Abstract / FREE Full Text 13. ↵ Gomez-Roldan , V. , Fermas , S. , Brewer , P.B. , et al. ( 2008 ) Strigolactone inhibition of shoot branching . Nature , 455 , 189 – 194 . OpenUrl CrossRef PubMed Web of Science 14. ↵ Hamiaux , C. , Drummond , R.S.M. , Janssen , B.J. , Ledger , S.E. , Cooney , J.M. , Newcomb , R.D. and Snowden , K.C . ( 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 . OpenUrl CrossRef PubMed 15. ↵ Hamiaux , C. , Drummond , R.S.M. , Luo , Z. , Lee , H.W. , Sharma , P. , Janssen , B.J. , Perry , N.B. , Denny , W.A. and Snowden , K.C . ( 2018 ) Inhibition of strigolactone receptors by N-phenylanthranilic acid derivatives: Structural and functional insights . J. Biol. Chem ., 293 , 6530 – 6543 . OpenUrl Abstract / FREE Full Text 16. ↵ Hassig , C.A. , Zeng , F.-Y. , Kung , P. , et al. ( 2014 ) Ultra-high-throughput screening of natural product extracts to identify proapoptotic inhibitors of Bcl-2 family proteins . J. Biomol. Screen ., 19 , 1201 – 1211 . OpenUrl CrossRef PubMed 17. ↵ 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 . OpenUrl CrossRef PubMed Web of Science 18. ↵ Jiang , L. , Liu , X. , Xiong , G. , et al. ( 2013 ) DWARF 53 acts as a repressor of strigolactone signalling in rice . Nature , 504 , 401 – 405 . OpenUrl CrossRef PubMed Web of Science 19. ↵ Kawada , K. , Takahashi , I. , Takei , S. , Nomura , A. , Seto , Y. , Fukui , K. and Asami , T . ( 2024 ) The evaluation of debranone series strigolactone agonists for germination stimulants in Orobanche species . J. Agric. Food Chem ., 72 , 19517 – 19525 . OpenUrl 20. ↵ Keppler , A. , Gendreizig , S. , Gronemeyer , T. , Pick , H. , Vogel , H. and Johnsson , K . ( 2003 ) A general method for the covalent labeling of fusion proteins with small molecules in vivo . Nat. Biotechnol ., 21 , 86 – 89 . OpenUrl CrossRef PubMed Web of Science 21. ↵ Khosla , A. , Morffy , N. , Li , Q. , et al. ( 2020 ) Structure-Function Analysis of SMAX1 Reveals Domains That Mediate Its Karrikin-Induced Proteolysis and Interaction with the Receptor KAI2 . Plant Cell , 32 , 2639 – 2659 . OpenUrl Abstract / FREE Full Text 22. ↵ Liu , Q. , Zhang , Y. , Matusova , R. , et al. ( 2014 ) Striga hermonthica MAX2 restores branching but not the Very Low Fluence Response in the Arabidopsis thaliana max2 mutant . New Phytol ., 202 , 531 – 541 . OpenUrl CrossRef PubMed Web of Science 23. ↵ Nelson , D.C . ( 2021 ) The mechanism of host-induced germination in root parasitic plants . Plant Physiol ., 185 , 1353 – 1373 . OpenUrl CrossRef PubMed 24. ↵ Ramil , C.P. , Dong , M. , An , P. , Lewandowski , T.M. , Yu , Z. , Miller , L.J. and Lin , Q . ( 2017 ) Spirohexene-tetrazine ligation enables bioorthogonal labeling of class B G protein-coupled receptors in live cells . J. Am. Chem. Soc ., 139 , 13376 – 13386 . OpenUrl CrossRef PubMed 25. ↵ Ritz , C. , Baty , F. , Streibig , J.C. and Gerhard , D . ( 2015 ) Dose-Response Analysis Using R . PLoS One , 10 , e0146021 . OpenUrl CrossRef PubMed 26. ↵ Sainsbury , F. , Thuenemann , E.C. and Lomonossoff , G.P . ( 2009 ) pEAQ: versatile expression vectors for easy and quick transient expression of heterologous proteins in plants . Plant Biotechnol. J ., 7 , 682 – 693 . OpenUrl CrossRef PubMed Web of Science 27. ↵ Santner , A. , Calderon-Villalobos , L.I.A. and Estelle , M . ( 2009 ) Plant hormones are versatile chemical regulators of plant growth . Nat. Chem. Biol ., 5 , 301 – 307 . OpenUrl CrossRef PubMed Web of Science 28. ↵ Scaffidi , A. , Waters , M.T. , Sun , Y.K. , Skelton , B.W. , Dixon , K.W. , Ghisalberti , E.L. , Flematti , G.R. and Smith , S.M . ( 2014 ) Strigolactone hormones and their stereoisomers signal through two related receptor proteins to induce different physiological responses in Arabidopsis . Plant Physiol ., 165 , 1221 – 1232 . OpenUrl Abstract / FREE Full Text 29. ↵ Seto , Y . ( 2023 ) Latest knowledge on strigolactone biosynthesis and perception . Biosci. Biotechnol. Biochem ., 88 , 1 – 7 . OpenUrl CrossRef 30. ↵ Seto , Y. , Yasui , R. , Kameoka , H. , et al. ( 2019 ) Strigolactone perception and deactivation by a hydrolase receptor DWARF14 . Nat. Commun ., 10 , 191 . OpenUrl CrossRef PubMed 31. ↵ Shabek , N. , Ticchiarelli , F. , Mao , H. , Hinds , T.R. , Leyser , O. and Zheng , N . ( 2018 ) Structural plasticity of D3-D14 ubiquitin ligase in strigolactone signalling . Nature , 563 , 652 – 656 . OpenUrl CrossRef PubMed 32. ↵ Sheard , L.B. , Tan , X. , Mao , H. , et al. ( 2010 ) Jasmonate perception by inositol-phosphate-potentiated COI1-JAZ co-receptor . Nature , 468 , 400 – 405 . OpenUrl CrossRef PubMed Web of Science 33. ↵ Soundappan , I. , Bennett , T. , Morffy , N. , Liang , Y. , Stanga , J.P. , Abbas , A. , Leyser , O. and Nelson , D.C . ( 2015 ) SMAX1-LIKE/D53 Family Members Enable Distinct MAX2-Dependent Responses to Strigolactones and Karrikins in Arabidopsis . Plant Cell , 27 , 3143 – 3159 . OpenUrl Abstract / FREE Full Text 34. ↵ Stanga , J.P. , Smith , S.M. , Briggs , W.R. and Nelson , D.C . ( 2013 ) SUPPRESSOR OF MORE AXILLARY GROWTH2 1 controls seed germination and seedling development in Arabidopsis . Plant Physiol ., 163 , 318 – 330 . OpenUrl Abstract / FREE Full Text 35. ↵ Takei , S. , Otani , M. , Ishikawa , T. , Suzuki , T. , Okabe , S. , Nishiyama , K. , Kawakami, N. and Seto, Y. ( 2024 ) Highly sensitive strigolactone perception by a divergent clade KAI2 receptor in a facultative root parasitic plant, Phtheirospermum japonicum . Plant Cell Physiol ., 65 , 1958 – 1968 . OpenUrl PubMed 36. ↵ Takei , S. , Uchiyama , Y. , Bürger , M. , Suzuki , T. , Okabe , S. , Chory , J. and Seto , Y . ( 2023 ) A Divergent Clade KAI2 Protein in the Root Parasitic Plant Orobanche minor Is a Highly Sensitive Strigolactone Receptor and Is Involved in the Perception of Sesquiterpene Lactones . Plant Cell Physiol ., 64 , 996 – 1007 . OpenUrl PubMed 37. ↵ Takeuchi , J. , Fukui , K. , Seto , Y. , Takaoka , Y. and Okamoto , M . ( 2021 ) Ligand-receptor interactions in plant hormone signaling . Plant J ., 105 , 290 – 306 . OpenUrl PubMed 38. ↵ Tal , L. , Palayam , M. , Ron , M. , Young , A. , Britt , A. and Shabek , N . ( 2022 ) A conformational switch in the SCF-D3/MAX2 ubiquitin ligase facilitates strigolactone signalling . Nat Plants , 8 , 561 – 573 . OpenUrl PubMed 39. ↵ Tan , X. , Calderon-Villalobos , L.I.A. , Sharon , M. , Zheng , C. , Robinson , C.V. , Estelle, M. and Zheng, N. ( 2007 ) Mechanism of auxin perception by the TIR1 ubiquitin ligase . Nature , 446 , 640 – 645 . OpenUrl CrossRef PubMed Web of Science 40. ↵ Toh , S. , Holbrook-Smith , D. , Stogios , P.J. , Onopriyenko , O. , Lumba , S. , Tsuchiya , Y. , Savchenko , A. and McCourt , P . ( 2015 ) Structure-function analysis identifies highly sensitive strigolactone receptors in Striga . Science , 350 , 203 – 207 . OpenUrl Abstract / FREE Full Text 41. ↵ Tsuchiya , Y. , Yoshimura , M. , Sato , Y. , et al. ( 2015 ) PARASITIC PLANTS. Probing strigolactone receptors in Striga hermonthica with fluorescence . Science , 349 , 864 – 868 . OpenUrl Abstract / FREE Full Text 42. ↵ Tsuzuki , K. , Suzuki , T. , Kuruma , M. , Nishiyama , K. , Hayashi , K.-I. , Hagihara , S. and Seto , Y . ( 2024 ) Radicle growth regulation of root parasitic plants by auxin-related compounds . Plant Cell Physiol ., 65 , 1377 – 1387 . OpenUrl PubMed 43. ↵ Umehara , M. , Cao , M. , Akiyama , K. , et al. ( 2015 ) Structural Requirements of Strigolactones for Shoot Branching Inhibition in Rice and Arabidopsis . Plant Cell Physiol ., 56 , 1059 – 1072 . OpenUrl CrossRef PubMed 44. ↵ Umehara , M. , Hanada , A. , Yoshida , S. , et al. ( 2008 ) Inhibition of shoot branching by new terpenoid plant hormones . Nature , 455 , 195 – 200 . OpenUrl CrossRef PubMed Web of Science 45. ↵ Uraguchi , D. , Kuwata , K. , Hijikata , Y. , et al. ( 2018 ) A femtomolar-range suicide germination stimulant for the parasitic plant Striga hermonthica . Science , 362 , 1301 – 1305 . OpenUrl Abstract / FREE Full Text 46. ↵ Wang , D. , Pang , Z. , Yu , H. , et al. ( 2022 ) Probing strigolactone perception mechanisms with rationally designed small-molecule agonists stimulating germination of root parasitic weeds . Nat. Commun ., 13 , 3987 . OpenUrl PubMed 47. ↵ Wang , L. , Wang , B. , Jiang , L. , 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 . OpenUrl Abstract / FREE Full Text 48. ↵ Wang , Y. , Yao , R. , Du , X. , Guo , L. , Chen , L. , Xie , D. and Smith , S.M . ( 2021 ) Molecular basis for high ligand sensitivity and selectivity of strigolactone receptors in Striga . Plant Physiol ., 185 , 1411 – 1428 . OpenUrl CrossRef PubMed 49. Waters , M.T. and Nelson , D.C . ( 2023 ) Karrikin perception and signalling . New Phytol ., 237 , 1525 – 1541 . OpenUrl PubMed 50. ↵ Waters , M.T. , Scaffidi , A. , Moulin , S.L.Y. , Sun , Y.K. , Flematti , G.R. and Smith , S.M . ( 2015 ) A Selaginella moellendorffii Ortholog of KARRIKIN INSENSITIVE2 Functions in Arabidopsis Development but Cannot Mediate Responses to Karrikins or Strigolactones . Plant Cell , 27 , 1925 – 1944 . OpenUrl Abstract / FREE Full Text 51. ↵ Wickham , H. ( 2016 ) Ggplot2: Elegant graphics for data analysis 2nd ed., Cham, Switzerland: Springer International Publishing . 52. ↵ Yao , R. , Ming , Z. , Yan , L. , et al. ( 2016 ) DWARF14 is a non-canonical hormone receptor for strigolactone . Nature , 536 , 469 – 473 . OpenUrl CrossRef PubMed 53. ↵ Yao , R. , Wang , F. , Ming , Z. , Du , X. , Chen , L. , Wang , Y. , Zhang , W. , Deng , H. and Xie , D . ( 2017 ) ShHTL7 is a non-canonical receptor for strigolactones in root parasitic weeds . Cell Res ., 27 , 838 – 841 . OpenUrl CrossRef PubMed 54. ↵ Yasui , R. , Seto , Y. , Ito , S. , Kawada , K. , Itto-Nakama , K. , Mashiguchi , K. and Yamaguchi , S . ( 2019 ) Chemical screening of novel strigolactone agonists that specifically interact with DWARF14 protein . Bioorg. Med. Chem. Lett ., 29 , 938 – 942 . OpenUrl PubMed 55. ↵ Zhao , L.-H. , Zhou , X.E. , Yi , W. , et al. ( 2015 ) Destabilization of strigolactone receptor DWARF14 by binding of ligand and E3-ligase signaling effector DWARF3 . Cell Res ., 25 , 1219 – 1236 . OpenUrl CrossRef PubMed 56. ↵ Zhou , F. , Lin , Q. , Zhu , L. , et al. ( 2013 ) D14–SCFD3-dependent degradation of D53 regulates strigolactone signalling . Nature , 504 , 406 – 410 . OpenUrl CrossRef PubMed Web of Science View the discussion thread. Back to top Previous Next Posted November 18, 2025. Download PDF Supplementary Material Email Thank you for your interest in spreading the word about bioRxiv. NOTE: Your email address is requested solely to identify you as the sender of this article. Your Email * Your Name * Send To * Enter multiple addresses on separate lines or separate them with commas. You are going to email the following In Vitro Dynamic and Quantitative Monitoring of Strigolactone-signaling Complex Formation by Time-resolved FRET Message Subject (Your Name) has forwarded a page to you from bioRxiv Message Body (Your Name) thought you would like to see this page from the bioRxiv website. 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Share In Vitro Dynamic and Quantitative Monitoring of Strigolactone-signaling Complex Formation by Time-resolved FRET Taiki Suzuki , Kotaro Nishiyama , Yusuke Kato , Chihiro Shinkai , Tomoya Ishikawa , Jekson Robertlee , Michio Kuruma , Shinya Hagihara , Marco Bürger , Kosuke Fukui , Tadao Asami , Yoshiya Seto bioRxiv 2025.11.18.688998; doi: https://doi.org/10.1101/2025.11.18.688998 Share This Article: Copy Citation Tools In Vitro Dynamic and Quantitative Monitoring of Strigolactone-signaling Complex Formation by Time-resolved FRET Taiki Suzuki , Kotaro Nishiyama , Yusuke Kato , Chihiro Shinkai , Tomoya Ishikawa , Jekson Robertlee , Michio Kuruma , Shinya Hagihara , Marco Bürger , Kosuke Fukui , Tadao Asami , Yoshiya Seto bioRxiv 2025.11.18.688998; doi: https://doi.org/10.1101/2025.11.18.688998 Citation Manager Formats BibTeX Bookends EasyBib EndNote (tagged) EndNote 8 (xml) Medlars Mendeley Papers RefWorks Tagged Ref Manager RIS Zotero Tweet Widget Facebook Like Google Plus One Subject Area Plant Biology Subject Areas All Articles Animal Behavior and Cognition (7642) Biochemistry (17715) Bioengineering (13907) Bioinformatics (42003) Biophysics (21470) Cancer Biology (18624) Cell Biology (25533) Clinical Trials (138) Developmental Biology (13390) Ecology (19935) Epidemiology (2067) Evolutionary Biology (24356) Genetics (15617) Genomics (22529) Immunology (17753) Microbiology (40432) Molecular Biology (17200) Neuroscience (88681) Paleontology (667) Pathology (2840) Pharmacology and Toxicology (4828) Physiology (7653) Plant Biology (15171) Scientific Communication and Education (2046) Synthetic Biology (4304) Systems Biology (9826) Zoology (2271)