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Strigolactones, host recognition signals for root parasitic plants and arbuscular mycorrhizal fungi, from Fabaceae plants

Kaori Yoneyama1, Xiaonan Xie1, Hitoshi Sekimoto2, Yasutomo Takeuchi1, Shin Ogasawara3, Kohki Akiyama3, Hideo Hayashi3 and Koichi Yoneyama1

1Weed Science Center, Utsunomiya University, 350 Mine-machi, Utsunomiya 321-8505, Japan; 2Faculty of Agriculture, Utsunomiya University, 350

Mine-machi, Utsunomiya 321-8505, Japan; 3Graduate School of Life and Environmental Sciences, Osaka Prefecture University, 1-1 Gakuencho,

Naka-ku, Sakai, Osaka 599-8531, Japan

Summary

• Both root parasitic plants and arbuscular mycorrhizal (AM) fungi take advantageof strigolactones, released from plant roots as signal molecules in the initial commu-nication with host plants, in order to commence parasitism and mutualism, respectively.• In this study, strigolactones in root exudates from 12 Fabaceae plants, includinghydroponically grown white lupin (Lupinus albus), a nonhost of AM fungi, werecharacterized by comparing retention times of germination stimulants on reverse-phase high-performance liquid chromatography (HPLC) with those of standards andby using tandem mass spectrometry (LC/MS/MS).• All the plant species examined were found to exude known strigolactones, suchas orobanchol, orobanchyl acetate, and 5-deoxystrigol, suggesting that thesestrigolactones are widely distributed in the Fabaceae. It should be noted that eventhe nonmycotrophic L. albus exuded orobanchol, orobanchyl acetate, 5-deoxystrigol,and novel germination stimulants.• By contrast to the mycotrophic Fabaceae plant Trifolium pratense, in whichphosphorus deficiency promoted strigolactone exudation, neither phosphorus nornitrogen deficiency increased exudation of these strigolactones in L. albus. Therefore,the regulation of strigolactone production and/or exudation seems to be closelyrelated to the nutrient acquisition strategy of the plants.

Key words: arbuscular mycorrhizal (AM) fungi, Fabaceae, nutrient acquisitionstrategy, parasitic plants strigolactones.

New Phytologist (2008) 179: 484–494

© The Authors (2008). Journal compilation © New Phytologist (2008) doi: 10.1111/j.1469-8137.2008.02462.x

Author for correspondence:Koichi YoneyamaTel: +81 28 649 5152Fax: +81 28 649 5155Email: yoneyama@cc.utsunomiya-u.ac.jp

Received: 21 January 2008Accepted: 7 March 2008

Introduction

The root parasitic plants Orobanche and Striga spp. aredevastating pests in agricultural production throughout theworld (Joel et al., 2007). These root parasites depend on hostplants for nutrients and water and cannot survive withoutparasitizing hosts. Their tiny seeds contain limited resourcesso that the parasites must connect to hosts within a week ofgermination. The seeds of these parasites germinate only

when they perceive host-derived chemicals, termed‘germination stimulants’, released from plant roots. The firstdescribed germination stimulant for Striga, named strigol,was isolated from the root exudates of a false host, cotton(Gossypium hirsutum) (Cook et al., 1966, 1972), and lateridentified from genuine hosts, sorghum (Sorghum bicolor),maize (Zea mays), and proso millet (Pennisetum glaucum)(Siame et al., 1993). Subsequently, sorgolactone was isolatedfrom S. bicolor root exudates (Hauck et al., 1992). Alectrol

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was purified from cowpea (Vigna unguiculata) root exudates(Müller et al., 1992) and recently identified as orobanchylacetate (Xie et al., 2008a). The first described Orobanchegermination stimulant, orobanchol, was isolated by Yokotaet al. (1998) from red clover (Trifolium pratense) rootexudates. Recently, 2′-epiorobanchol and solanacol werecharacterized from root exudates of tobacco (Nicotiana tabacum),a host of Phelipanche ramosa (formally called Orobancheramosa) (Xie et al., 2007). In addition, two novel stimulants,sorgomol (Awad et al., 2006; Xie et al., 2008b) and a putativedidehydro-orobanchol (strigol) isomer (Sato et al., 2003;Xie et al., 2007), were identified in the root exudates ofseveral Poaceae species and the Solanaceae species N. tabacumand tomato (Solanum lycopersicum). The structure of thisdidehydro-orobanchol isomer has not yet been clarified.These strigol-related germination stimulants are collectivelycalled strigolactones (Fig. 1).

Among the strigolactones, 5-deoxystrigol was originallyisolated as a branching factor of arbuscular mycorrhizal (AM)fungi from root exudates of Lotus japonicus (Akiyama et al.,2005). We also identified 5-deoxystrigol as one of majorgermination stimulants of S. bicolor, Z. mays, and pearl millet(Pennisetum typhoideum) (Awad et al., 2006).

The AM fungi, which are obligate symbionts, are incapableof completing their life cycle without residing in their host

roots. After spore germination and hyphal growth, the hyphalbranching of AM fungi occurs in the vicinity of host roots.Because the phenomenon does not occur in the vicinity ofroots of nonhosts, including rapeseed (Brassica napus) andwhite lupin (Lupinus albus), hyphal branching is consideredto be the host recognition process (Giovannetti et al., 1993).In addition, strigolactones were found to induce a rapidincrease in mitochondrial density and changes in the shapeand movement of the organelles in AM fungi (Besserer et al.,2006). Such activation of the mitochondria might lead tothe oxidation of lipids, which are the main form of carbonstorage in AM fungal spores. Therefore, strigolactones maybe crucial components of root exudates that switch on lipidcatabolism at the presymbiotic stage of the fungus (Bessereret al., 2006; Akiyama, 2007). Furthermore, strigolactonesinduce gene expression of Gigaspora margarita CuZn super-oxide dismutase (GmarCuZnSOD) (Lanfranco et al., 2005)and chemotrophic growth of Glomus mosseae hyphae (Sbrana& Giovannetti, 2005). The AM association is by far the mostwidespread association between microorganisms and higherplants. Within the angiosperms, at least 80% of the species areable to form AM symbioses (Harrison, 2005). Therefore, ifstrigolactones are indispensable for host recognition of AMfungi, they may be widely distributed in the plant kingdom(Akiyama, 2007). However, characterizations of strigolactones

Fig. 1 Chemical structures of natural strigolactones and the synthetic analog GR24.

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have been conducted for only a few plant species as plantsexude trace amounts of unstable strigolactones. Accordingly,it is necessary to clarify the distribution of strigolactones in theplant kingdom to understand the chemical communicationsin the rhizosphere between plants and AM symbionts, andplants and root parasites.

We have developed a specific and rapid analytical methodfor known strigolactones using high-performance liquidchromatography (HPLC) connected to tandem massspectrometry (LC/MS/MS) (Sato et al., 2003). LC/MS/MSanalyses revealed the presence of 5-deoxystrigol in the rootexudates of S. bicolor, Z. mays, and P. typhoideum (Awad et al.,2006). Besserer et al. (2006) identified sorgolactone as abranching factor in S. bicolor root exudates using LC/MS/MS. Furthermore, using LC/MS/MS, we demonstrated thatnutrient deficiencies affect strigolactone exudation: in T. pratense,phosphorus (P) deficiency significantly promoted orobancholexudation (Yoneyama et al., 2007a), while in S. bicolor, nitrogen(N) deficiency as well as P deficiency enhanced 5-deoxystrigolexudation (Yoneyama et al., 2007b).

To date, we have examined a wide range of plant speciesincluding crops, weeds, and even trees for the production ofstrigolactones and have found that plants produce diversemixtures of known and unknown strigolactones. Althoughsome of these unknown strigolactones have been purified andsubjected to structural elucidation (Yokota et al., 1998; Xieet al., 2007, 2008a,b), at least several novel strigolactones remainto be characterized (K. Yoneyama, unpublished). Therefore,to compare strigolactone production among different plantspecies, the major strigolactones in each plant species shouldfirst be identified. As in the case of the Poaceae species (Awadet al., 2006), the major strigolactones of plant species withinthe same family are expected to be similar. We thus focused onthe Fabaceae for the comparison of strigolactone productionwithin the family as we had already identified major strigolac-tones produced by T. pratense (Yokota et al., 1998).

In this paper we extended the characterization of strigolac-tones in root exudates of 12 Fabaceae plant species, includingL. albus, a nonhost of AM fungi, by comparing retention timesof germination stimulants on reverse-phase (RP)-HPLC withthose of standards and by using LC/MS/MS. In addition, theeffects of N deficiency and P deficiency on strigolactoneexudation by L. albus were examined to clarify whether theregulation of strigolactone exudation is related to the nutrientacquisition strategy of plants.

Materials and Methods

Chemicals

(+)-Orobanchol was a generous gift of Emeritus Prof. KenjiMori (The University of Tokyo, Tokyo, Japan). GR24 waskindly provided by Prof. Binne Zwanenburg (RadboudUniversity, Nijmegen, the Netherlands). (+)-Orobanchyl acetate

was purified from root exudates of T. pratense (Xie et al.,2008a). (+)-Solanacol and the didehydro-orobanchol isomerwere purified from root exudates of N. tabacum (Xie et al.,2007). (±)-5-Deoxystrigol was prepared as reported previously(Akiyama et al., 2005). The other chemicals of analyticalgrade and HPLC solvents were obtained from Kanto ChemicalCo. Ltd (Tokyo, Japan) and Wako Pure Chemical IndustriesLtd (Osaka, Japan).

Plant material

Orobanche minor Sm. seeds were collected from mature plantsthat were parasites of Trifolium pratense L. grown in theWatarase basin of Tochigi Prefecture, Japan. Seeds of Glycinemax (L.) Merrill, Phaseolus vulgaris L., Vicia faba L., Arachishypogaea L., Astragalus sinicus L., Medicago sativa L., Pisumsativum L., Trifolium incarnatum L., Vigna angularis (Willd.)Ohwi & Ohashi, and Psophocarpus tetragonolobus L. wereobtained from a local supplier. Seeds of Cicer arietinum L.,and Lupinus albus L. were generously supplied by Dr YaakovGoldwasser (Hebrew University of Jerusalem, Jerusalem,Israel) and Dr Jun Wasaki (Hiroshima University, Hiroshima,Japan), respectively.

Growth conditions and collection of root exudates

Plant seeds were surface-sterilized in 70% ethanol for 2 minand then 1% NaClO for 2 min. After thoroughly rinsingwith sterile distilled water, the seeds were germinated onmoistened filter paper in a container at 23°C (L. albus andC. arietinum) or 25°C (the remainder) in the dark for 3 d.Seedlings (n = 20–100) were transferred to a strainer(28 × 23 × 9 cm, width × length × height (W × L × H)) linedwith a sheet of gauze moistened by placing it in a slightlylarger container (28.5 × 23.5 × 11 cm, W × L × H) containing1 l of tap water as the culture medium in a growth chamberwith a 14 : 10 h photoperiod at 120 µmol photons m–2 s–1

at 23 : 20°C (L. albus and C. arietinum) or 25 : 23°C (theremainder). The plants were grown in tap water for 7 dand then transferred to 1/2 Tadano and Tanaka medium(Tadano & Tanaka, 1980) without P and grown for another3 d. Two strainers were used for each plant species. Then,the two strainers were transferred to a larger container(53.5 × 33.5 × 14 cm, W × L × H) containing 10 l of tapwater and 1 mM CaCl2. Root exudates released into culturemedium were adsorbed by activated charcoal with usingcirculation pumps (Akiyama et al., 2005). The plants weregrown for 8 d, during which the culture medium andactivated charcoal were exchanged every other day.

Extraction of root exudates

Root exudates adsorbed on the charcoal were eluted withacetone. After the acetone was evaporated in vacuo, the

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residue was dissolved in 50 ml of water and extracted 3 timeswith 50 ml of ethyl acetate. The ethyl acetate extracts werecombined, washed with 0.2 M K2HPO4 (pH 8.3), dried overanhydrous MgSO4, and concentrated in vacuo. These crudeextracts were stored in sealed glass vials at 4°C until use.

Characterization of strigolactones

Characterization of strigolactones in the root exudates wasconducted by comparing retention times of germinationstimulants on RP-HPLC with those of synthetic (or natural)standards and by using LC/MS/MS (Sato et al., 2003, 2005;Awad et al., 2006).

HPLC separation

HPLC separation was conducted with a U980 HPLCinstrument (Jasco, Tokyo, Japan) fitted with an ODS (C18)column (Mightysil RP-18, 2 × 250 nm, 5 µm; Kanto ChemicalsCo., Ltd., Tokyo, Japan). The crude extracts were dissolved in60% methanol and filtered through spin columns (Ultra-FreeMC, 0.45 µm pore size; Milipore, Tokyo, Japan), and 10 µlwas injected. The mobile phase was 60% methanol in waterand was changed to 100% methanol 30 min after injection.The column was then washed with 100% methanol for20 min. The flow rate was 0.2 ml min–1 and the columntemperature was set to 40°C.

Mass spectrometry

Mass spectrometry was performed with a Quattro LC massspectrometer (Micromass, Manchester, UK) equipped withan electrospray source. The drying and nebulizing gas wasnitrogen generated from pressurized air in an N2G nitrogengenerator (Parker-Hanifin Japan, Tokyo, Japan). The nebulizergas flow was set to approx. 100 l h–1, and the desolvation gasflow to 500 l h–1. The interface temperature was set to 400°C,and the source temperature to 150°C. The capillary and conevoltages were adjusted to orobanchol and to the positiveionization mode. MS/MS experiments were conducted usingargon as the collision gas and the collision energy was set to16 eV. The collision gas pressure was 0.15 Pa. For thedetection of known strigolactones, we used six-channelmultiple reaction monitoring (MRM). The six transitions ofm/z 339 > 242, 353 > 256, 365 > 268, 367 > 270, 369 > 272,and 411 > 254 were monitored for sorgolactone, 5-deoxystrigol, solanacol, didehydro-orobanchol (strigol), strigol(orobanchol and sorgomol), and orobanchyl acetate (strigylacetate), respectively. Data acquisition and analysis wereperformed with the MassLynx software (ver. 4.1).Quantification of strigolactones was carried out usingsynthetic or natural standards in a manner similar to thatfor orobanchol, strigol, and 5-deoxystrigol (Sato et al., 2003,2005; Yoneyama et al., 2007a,b).

Germination assays

A portion of the ethyl acetate extracts dissolved in 60%methanol was fractionated by RP-HPLC operated under thesame conditions as for LC/MS/MS analyses and the fractionscollected every minute were examined for O. minor seedgermination stimulation (Goldwasser et al., 2008). For this,preliminary germination assays were conducted to determinean optimal dilution of each crude extract, because, when anexcessive amount of crude extract was loaded for HPLC,germination stimulation activity was distributed broadlyin many fractions. The amounts of crude extracts used forgermination assays were 1/1000 that used for LC/MS/MSanalyses for P. tetragonolobus, 1/500 for A. hypogaea andV. faba, and 1/100 for the other species.

Germination assays on O. minor seeds were conductedas reported previously (Yoneyama et al., 2007a). The surface-sterilized O. minor seeds, c. 20 each, were placed on 6-mmglass fiber discs (Whatman GF/A) and c. 90 discs wereincubated in a 9-cm sterile Petri dish lined with a sheet offilter paper (No. 2, Advantec, Tokyo, Japan) and wetted with6 ml of sterile Milli-Q water in the dark at 23°C for 7 d as a‘conditioning period’ during which the seeds become respon-sive to germination stimulants. Then, four discs carrying theconditioned seeds were transferred to a 5-cm sterile Petri dishprepared as follows. An aliquot of root exudate samples in 60%methanol or methanol solutions of authentic strigolactoneswas added to a 5-cm Petri dish lined with a filter paper. Thesolvent was allowed to evaporate before the discs carrying theconditioned seeds were placed on the filter paper and treatedwith sterile Milli-Q water (650 µl). The Petri dishes weresealed, enclosed in polyethylene bags, and placed in thedark at 23°C for 4–5 d. Seeds treated with or without GR24(10–6 M) were always included as positive and negative controls.Seeds were considered germinated when the radicle protrudedthrough the seed coat.

Hyphal branching assay

Hyphal branching assays for Gigaspora margarita Becker &Hall were conducted as reported previously (Akiyama et al.,2005). Spores of G. margarita (CGC1411; Central Glass Co.,Tokyo, Japan), surface-sterilized with 0.2% NaClO and 0.05%Triton X-100, were inserted into a 0.2% Phytagel gel (Sigma-Aldrich, Tokyo, Japan) containing 3 mM MgSO4 in 60-mmplastic Petri dishes. The dishes were incubated vertically for5–7 d in a 2% CO2 incubator at 32°C. Secondary hyphaeemerging from a primary hypha, which grew upward in anegative geotropic manner in the gel, were used for assay. Testsamples were first dissolved in acetone and then diluted with70% ethanol in water. Paper discs (6 mm) loaded with 15 µlof test sample solution were placed in front of the tips of thesecondary hyphae. The control was 70% ethanol in water. Thehyphae branch patterns were observed 24 h after treatment.

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The sample was scored as positive for hyphal branching if newhyphal branches formed from the treated secondary hyphaeor primary hyphae located proximal to the paper discs. Theassay was repeated at least twice, using between three and fivedishes for each concentration.

The clusters of hyphal branches consisting of higher ordersof hyphae (third, fourth, and fifth hyphae) are induced fromthe secondary hyphae by treatment with positive samplessuch as root exudates and pure strigolactones. In the controltreatment (70% ethanol in water), no hyphae or an occasionalsingle branching hypha is induced from the treated secondaryhyphae located proximal to the paper discs. This distinctdifference in hyphal morphology enables us to distinguishpositive samples from negative controls qualitatively.

Effects of nutrient deficiencies on strigolactone exudation by L. albus

After germination, 20 seedlings of L. albus were grown in tapwater for 10 d and then subjected to each nutrient condition.Half-strength Tadano and Tanaka media (Tadano & Tanaka,1980) was used as the control medium containing 2.43 mMN and 160 µM P. Low-N and low-P nutrient media had 120 µMN and 8 µM P, respectively. After 10 d of acclimatization inthe test media, the growth media containing root exudates (pluswashings) collected daily (1.5 l) were extracted with ethylacetate as reported previously (Yoneyama et al., 2007a,b).

Results

Characterization of strigolactones in root exudates of Fabaceae plants

For the characterization of strigolactones produced by theFabaceae species, we collected relatively large amounts of rootexudates as we did not know how much strigolactones theywould produce. The plants were grown in tap water and rootexudates were collected using activated charcoal, because theproduction of strigolactones seemed to be enhanced underthese conditions (Yoneyama et al., 2001; Akiyama et al., 2005).We first tried to examine qualitative but not quantitativedifferences in strigolactone production among plant species.In the germination assays, we used O. minor because this parasitecould be handled without any strict quarantine restrictionsand in addition O. minor has been reported to have a relativelybroad host range (Parker, 1986; Parker & Riches, 1993) andthus its seeds are expected to be sensitive to various strigo-lactones. Not all of the 12 Fabaceae species examined in thisstudy have been reported as hosts for Orobanche spp. Some ofthem are hosts of Orobanche crenata, Phelipanche ramosa andPhelipanche aegyptiaca (Parker, 1986; Parker & Riches, 1993).

In the preliminary germination and hyphal branchingassays with crude extracts, all of the 12 Fabaceae species werefound to exude germination stimulants of O. minor and

branching factors of G. margarita. Then, each crude extractwas examined for the presence of known strigolactones bycomparing retention times of germination stimulants onRP-HPLC with those of synthetic or natural standards andby using LC/MS/MS. The results clearly demonstrated thatall of the plants examined, including L. albus, a nonhost ofAM fungi, produced known strigolactones.

A four-channel MRM chromatogram and the distributionof germination stimulation activity on O. minor after RP-HPLC separation of L. albus root exudates are shown inFig. 2. Among known strigolactones, orobanchol, orobanchylacetate, and 5-deoxystrigol were identified in the transitionsof m/z 369 > 272, 411 > 254, and 353 > 256 at retention timesof 7.9, 16.4 and 27.3 min, respectively, by LC/MS/MS (Fig. 2a).These assignments were confirmed by co-chromatographywith synthetic and natural standards (data not shown). Ger-mination stimulation activities on O. minor were associatedwith the fractions whose retention times corresponded tothose of orobanchol (fraction 8), orobanchyl acetate (fractions15 and 16), and 5-deoxystrigol (fraction 28) (Fig. 2b). Inaddition to these known strigolactones, a distinct peak wasdetected in the MRM channel monitoring the transition ofm/z 367 > 270 at a retention time of 6.6 min, and the activitywas observed in fraction 6 corresponding to this peak in theMRM chromatogram. This was identified as a didehydro-orobanchol isomer which had been detected in the rootexudates of the Solanaceae plants N. tabacum (Xie et al.,2007) and S. lycopersicum (Sato et al., 2003). This strigolac-tone was the third, minor stimulant produced by T. pratense(Yokota et al., 1998) but its structure has not yet beenelucidated. Furthermore, there was a distinct germinationstimulation activity eluted in fraction 11 (11–12 min), whereasthere were no peaks at the corresponding retention time inthe six-channel MRM chromatogram, indicating the presenceof an unknown germination stimulant. Accordingly, L. albuswas found to exude the didehydro-orobanchol isomer, orob-anchol, orobanchyl acetate, 5-deoxystrigol and an unknowngermination stimulant (Fig. 2a,b).

Figure 3 shows a four-channel MRM chromatogram andthe distribution of germination stimulation activity on O. minorafter RP-HPLC separation of P. sativum root exudates. Orob-anchol, orobanchyl acetate, 5-deoxystrigol, and the didehydro-orobanchol isomer were detected (Fig. 3a) by LC/MS/MSand their identities were confirmed by co-chromatographywith strigolactone standards (data not shown). However,germination stimulation activity was not observed in thefractions corresponding to the retention times of thedidehydro-orobanchol isomer, orobanchol, and 5-deoxystrigol(Fig. 3b). Fraction 16 (16–17 min), corresponding to theretention time of orobanchyl acetate, showed weak germinationstimulation. Instead, potent germination stimulation activitywas observed in fraction 12 (12–13 min), indicating that themajor germination stimulant of P. sativum was an unknowncompound.

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Distinct germination stimulant activity in fraction 12 afterRP-HPLC separation was detected not only in the rootexudates of P. sativum but also in those of A. sinicus,A. hypogaea, C. arietinum, M. sativa, T. incarnatum, and V. faba.Therefore, this novel stimulant seems to be distributed widelyin the Fabaceae. However, it remains unclear whether theseplants exude the same compound.

Table 1 summarizes the distribution of strigolactones amongthe Fabaceae plants examined. In this table, strigolactonesidentified by LC/MS/MS are designated as ‘MS’ and ‘G’ meansthat germination stimulation activity was detected in thefraction corresponding to the retention time of the strigolactonestandard. It should be noted that germination assays wereconducted at a particular dilution of the root exudates atwhich major stimulants were clearly separated by RP-HPLC.Therefore, the lack of G in Table 1 means that the contributionof the corresponding strigolactone to the overall germinationstimulation activity of the root exudates was rather small.

Orobanchyl acetate was detected in the root exudates of allthe Fabaceae plants examined. Orobanchol and 5-deoxystrigolwere present in the root exudates of 11 of the 12 Fabaceaeplants. Therefore, these three strigolactones, orobanchol,orobanchyl acetate, and 5-deoxystrigol, were found to bemajor strigolactones in the Fabaceae plants. In addition tothese major strigolactones, the didehydro-orobanchol isomerwas detected by LC/MS/MS and its activity was confirmedby germination assays in the root exudates of A. sinicus,C. arietinum, and L. albus. Solanacol was identified only inthe root exudates of T. incarnatum. Although 5-deoxystrigolwas detected by LC/MS/MS in the root exudates of G. max,P. vulgaris, P. sativum, and V. faba, the RP-HPLC fractionscorresponding to the retention time of 5-deoxystrigol wereinactive in the germination assays (Table 1), indicating that5-deoxystrigol was a minor germination stimulant in thesespecies. None of these Fabaceae plants produced detectablelevels of strigol, sorgolactone, or sorgomol.

Fig. 2 (a) Four-channel multiple reaction monitoring (MRM) chromatogram of the root exudates from Lupinus albus, a nonhost of arbuscular mycorrhizal (AM) fungi, where the transitions of m/z 367 > 270, 369 > 272, 411 > 254, and 353 > 256 were monitored for didehydro-orobanchol, orobanchol, orobanchyl acetate, and 5-deoxystrigol (and their isomers), respectively. (b) Distribution of germination stimulation activity on Orobanche minor after reverse-phase high-performance liquid chromatography (HPLC) separation of the root exudates. All the fractions were tested for germination stimulation activity but only the numbers of active fractions are presented.

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Fig. 3 (a) Four-channel multiple reaction monitoring (MRM) chromatogram of root exudates from Pisum sativum. The channels were set for the same transitions as in Fig. 2a. (b) Distribution of germination stimulation activity on Orobanche minor after reverse-phase high-performance liquid chromatography (HPLC) separation of the root exudates. All the fractions were tested for germination stimulation activity but only the numbers of active fractions are presented.

Table 1 Distribution of strigolactones in Fabaceae plants

Scientific name Solanacol Didehydro-orobanchol Orobanchol Orobanchyl acetate 5-Deoxystrigol

Arachis hypogaea – – MS/G MS/G MS/GAstragalus sinicus – MS/G MS/G MS/G MS/GCicer arietinum – MS/G – MS/G MS/GGlycine max – – MS/G MS/G MSLupinus albus – MS/G MS/G MS/G MS/GMedicago sativa – – MS/G MS/G MS/GPhaseolus vulgaris – – MS/G MS/G MSPisum sativum – MS MS MS/G MSPsophocarpus tetragonolobus – – MS/G MS/G MS/GTrifolium incarnatum MS/G – MS/G MS/G –Vicia faba – – MS/G MS/G MSVigna angularis – – MS/G MS/G MS/G

Strigolactones were characterized by using tandem mass spectrometry (LC/MS/MS) and by comparing retention times of germination stimulation activity on reverse-phase high-performance liquid chromatography (HPLC) with those of natural and synthetic standards. MS indicates that the strigolactone was detected by LC/MS/MS. G indicates that germination stimulation activity on Orobanche minor seeds was observed at the retention time corresponding to that of the strigolactone. It should be noted that only major germination stimulants could be detected in the germination assays.

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Effects of N and P deficiencies on strigolactone exudation by L. albus

The nonmycotrophic L. albus species was found to exude thedidehydro-orobanchol isomer, orobanchol, orobanchylacetate, 5-deoxystrigol, and an unknown stimulant (Fig. 2a,b).Among the strigolactones present in the root exudates ofL. albus, orobanchol, orobanchyl acetate and 5-deoxystrigolcould be quantified by LC/MS/MS. Therefore, the effects of Pand N deficiencies on the exudation of orobanchol, orobanchylacetate, and 5-deoxystrigol by L. albus were examined.

The P contents of shoots and roots of L. albus plants grownunder low-P conditions were 2.6 and 3.2 mg (g DW)–1,which were 40% and 20% those of the controls, respectively,and thus the plants were in a state of P deficiency. Similarly,the N contents of the plant material subjected to N deficiencywere < 50% those of the controls.

In the case of the mycotrophic Fabaceae plant T. pratense, areduced supply of P but not of other minerals significantlyincreased the exudation of orobanchol by the roots (Yoneyamaet al., 2007a). In contrast, deficiency of neither P nor Npromoted the exudation of orobanchol, orobanchyl acetate,or 5-deoxystrigol by L. albus roots (Fig. 4a). In fact, both Nand P deficiencies tended to reduce exudation of these strig-olactones. The nonmycotrophic L. albus grown hydroponicallyunder P deficiency released 1.5 ± 0.5 pg g–1 root FW of orob-anchol and 37.5 ± 7.5 pg g–1 root FW of orobanchyl acetateover 3 d (mean ± SE, n = 3) as shown in Fig. 4a. By contrast,under the same growth conditions, the mycotrophic T. pratenseexuded 36 000 ± 4200 pg g–1 root FW of orobanchol and153 000 ± 19 700 pg g–1 root FW of orobanchyl acetate over3 d (mean ± SE, n = 3) (Yoneyama et al., 2007a).

Lupinus albus exudes the didehydro-orobanchol and anunknown germination stimulant in addition to these strigol-actones, as shown in Fig. 2(a,b), and thus deficiency of N and/or P may affect exudation of those stimulants which cannotbe quantified as their pure standards are not yet available.The amounts of the pure didehydro-orobanchol isomerisolated from N. tabacum root exudates were too small toweigh accurately. Therefore, crude extracts of root exudatesfrom the plants grown under P or N deficiency were examinedfor their germination stimulation on O. minor seeds (Fig. 4b).

A 4.5-ml equivalent of the control culture medium inducedc. 50% germination, while those of N and P deficiencies atthe same dosage elicited < 10% germination (Fig. 4b). At45- and 450-ml equivalents of culture medium, there wereno significant differences in the germination stimulationactivities among the root exudates from the plants grownunder the different nutrient conditions. The reduction ingermination rate at the highest dosage of root exudate fromthe control was attributable to germination inhibitors producedsimultaneously. The germination stimulation activities in theculture media therefore seemed to be nearly proportionalto the amounts of orobanchol, orobanchyl acetate, and 5-

deoxystrigol, suggesting that the exudation of the didehydro-orobanchol isomer and the unknown germination stimulantappeared to be affected similarly to the other strigolactones.

Discussion

In the present study, strigolactones exuded from 12 Fabaceaeplants were characterized by an RP-HPLC separation–germination assay and by LC/MS/MS. Orobanchyl acetate

Fig. 4 (a) Exudation of orobanchol (white bars), orobanchyl acetate (gray bars), and 5-deoxystrigol (dark gray bars) by Lupinus albus roots and (b) germination stimulation activity of L. albus root exudates from plants grown under nutrient deficiencies in hydroponic culture medium over 3 d. The control medium (white circles) contained 160 µM phosphorus (P) and 2.43 mM nitrogen (N) which were reduced to 1/20 in the low-P (8 µM P; triangles) and low-N (120 µM N; black circles) media. (a) Root exudate samples collected daily were analyzed separately by tandem mass spectrometry (LC/MS/MS). The data are sums of exudations over 3 d. The experiments were repeated three times. Values represent the means ± SE. (b) Root exudate samples collected daily for 3 d were combined and used in germination assays with Orobanche minor seeds. Aliquots of methanol solutions of the root exudate samples corresponding to 4.5, 45, and 450 ml of medium were transferred to 5-cm Petri dishes lined with a filter paper, and the solvent was allowed to evaporate before the discs carrying the conditioned O. minor seeds were placed on the filter paper and treated with distilled water (650 µl). The experiments were repeated three times. Values represent the means ± SE.

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(alectrol) was detected in root exudates from all of these plants,and most of them exuded orobanchol and 5-deoxystrigol.Therefore, orobanchol, orobanchyl acetate, and 5-deoxystrigolappear to be major germination stimulants in the Fabaceae.In addition to these strigolactones, a didehydro-orobancholisomer and solanacol were identified in the root exudatesfrom four and one species, respectively. Furthermore, twonovel stimulants, one from P. sativum and the other fromL. albus root exudates, were detected. The stimulant found inthe root exudate of P. sativum seems to be present in the rootexudates of A. sinicus, A. hypogaea, C. arietinum, M. sativa,and V. faba. After several steps of purification, this stimulantappeared to be a novel strigolactone with a molecular weightof 404 Da, while its structure has not yet been fully elucidatedas the amounts purified from P. sativum root exudates werenot sufficient for comprehensive spectroscopic analyses.

In the case of P. sativum, although distinct peaks of thedidehydro-orobanchol, orobanchol, orobanchyl acetate, and5-deoxystrigol were observed in the MRM chromatogram(Fig. 3a), germination stimulation activities were not detectedin the RP-HPLC fractions corresponding to the retentiontimes of orobanchol and 5-deoxystrigol. Only a weak ger-mination stimulation activity was associated with fraction 16,corresponding to the retention time of orobanchyl acetate(Fig. 3b). This implies that these strigolactones contributedto the overall germination stimulation activity in the rootexudate only to a small extent as compared with the novelstrigolactone eluted in fractions 12 and 13 and orobanchylacetate in fraction 16.

In the LC/MS/MS analyses, each strigolactone was moni-tored with the MRM channel specific to it at an independentsensitivity. For example, in Fig. 3a, the ion intensity in thechannel for orobanchyl acetate (1.95 × 107) was 100-fold largerthan those for orobanchol (1.15 × 105) and 5-deoxystrigol(7.14 × 104). Given the fact that, in electrospray ionization(ESI) mass spectrometry, ion intensities are not always pro-portional to the concentrations of analytes, and the ionizationefficiency of orobanchol is c. 10-fold lower than those oforobanchyl acetate and 5-deoxystrigol, the amounts oforobanchyl acetate would be at least 10-fold larger than thoseof orobanchol and 5-deoxystrigol. We could not increaseamounts of root exudate samples for RP-HPLC separation,because the activity was distributed broadly in many fractionswhen an excessive amount of sample was loaded for HPLC.

Similar results were obtained with the root exudates ofG. max, P. vulgaris, and V. faba, where 5-deoxystrigol wasdetected by LC/MS/MS but germination stimulation activitywas not observed in the fraction(s) corresponding to the retentiontime of 5-deoxystrigol. In addition to the differences in thecontents of strigolactones as discussed above, the differencesin sensitivity of root parasite seeds to various strigolactonesand the presence of inhibitors also affected the results ofgermination assays. In fact, O. minor used in the germinationassays was c. 100-fold more sensitive to orobanchol than to

5-deoxystrigol; 0.1 nM orobanchol elicited c. 60% germinationand a similar level of germination was observed at 10 nM5-deoxystrigol.

Although many authors have suggested that strigolactonesare distributed widely in the plant kingdom, the isolation andidentification of strigolactones have been hampered by theextremely low concentrations produced and exuded by hostroots as well as their relative instability. The recent develop-ment of an analytical method using LC/MS/MS enablesidentification and quantification of known strigolactones (Satoet al., 2003, 2005; Awad et al., 2006; Besserer et al., 2006;Yoneyama et al., 2007a,b; Xie et al., 2007) and, in addition,the search for novel strigolactones (Awad et al., 2006; Xieet al., 2007). Here we demonstrated that the 12 Fabaceaeplants, including the nonmycotrophic L. albus, exude at leastthree strigolactones, and that, among the known strigolactones,orobanchol, orobanchyl acetate, and 5-deoxystrigol are themajor ones produced by these plants grown hydroponically.

It is intriguing that all the plants examined so far exudemixtures of at least two strigolactones. Although the questionof why plants produce and exude mixtures of strigolactoneshas not yet been answered, quantitative and/or qualitativedifferences in the strigolactone compositions may be one ofthe key factors determining the host specificity of AM fungiand root parasites. Therefore, the effects of various combina-tions of strigolactones on both hyphal branching of AM fungiand seed germination of root parasites need to be examined.

Orobanchol and orobanchyl acetate (alectrol) were firstisolated from root exudates of T. pratense and V. unguiculataas germination stimulants, respectively. Orobanchol was alsoidentified in root exudates from Solanaceae plants includingN. tabacum (Xie et al., 2007) and S. lycopersicum (Sato et al.,2003), and from the Compositae marigold (Targetes patula)(K. Yoneyama, unpublished). Orobanchyl acetate was detectedin several Compositae plants (K. Yoneyama, unpublished).However, these strigolactones have not been found in theroot exudates from any members of the Poaceae examined todate (Awad et al., 2006; K. Yoneyama, unpublished). Theseresults suggest that orobanchol and orobanchyl acetate seemto be distributed in dicotyledons but not in monocotyledons.

5-Deoxystrigol, originally isolated as a hyphal branchinginducer for AM fungi from the root exudate of Lotus japonicus(Akiyama et al., 2005), has been shown to be one of majorstrigolactones in monocotyledonous plants including S. bicolor,Z. mays and P. typhoideum (Awad et al., 2006). As all theFabaceae plants examined in this study, with the exception ofT. incarnatum, were found to exude 5-deoxystrigol, 5-deoxystrigolappears to be widely distributed in both monocotyledonsand dicotyledons. This is in good agreement with the pro-posed biosynthetic pathway for strigolactones, originatingfrom carotenoids, where 5-deoxystrigol serves as the precursorof all the other known strigolactones (Matusova et al., 2005;Bouwmeester et al., 2007). The oxidation of 5-deoxystrigolproduces mono-hydroxy-strigolactones such as orobanchol,

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strigol, and the recently identified sorgomol (Xie et al., 2008b).It is likely that sorgomol is then converted to sorgolactoneby subsequent oxidation and decarboxylation (Xie et al., 2008b;Rani et al., in press). Among the plants examined to date,T. incarnatum, N. tabacum (Xie et al., 2007) and S. lycopersicum(unpublished data) did not produce detectable amounts of5-deoxystrigol. It is interesting that these three plant speciesexuded solanacol, a strigolactone containing a benzene ring.

AM fungi supply mineral nutrients, especially P, to hostplants by extending beyond the depletion zone for P aroundthe root, the external mycelia improving P absorption. How-ever, AM fungi are absent under all environmental conditionsin the Brassicaceae and Chenopodiaceae, and are also quiterare or absent in many members of the Proteaceae and othertypical root cluster-forming plant species including L. albus(Marschner, 1993). The proximity of roots of nonhostsincluding L. albus did not elicit hyphal branching of AMfungi (Giovannetti et al., 1993). An early hypothesis thatnonhost plants do not induce the morphogenetical eventsuggested that roots of nonhosts secrete compounds into therhizosphere that inhibit AM colonization (Fontenla et al.,1999). Schreiner & Koide (1993) showed that members ofBrassicaceae have the potential to produce significant quantitiesof antifungal compounds in roots, probably isothiocyanates.An alternative explanation for the lack of AM colonization ofnonhosts is that nonhosts fail to produce branching factorsrequired by AM fungi for host recognition. In this study,it was demonstrated that the nonmycotrophic L. albus grownhydroponically also produces and exudes strigolactones, andtherefore the hypothesis that nonhosts of AM fungi fail toexude strigolactones may be excluded.

The amounts of strigolactones released from L. albus,however, were quite low compared with mycotrophic plants.Indeed, the amounts of orobanchol and orobanchyl acetate,per unit root fresh weight, released from L. albus grown underP starvation were only 1/24 000 and 1/4080 that fromT. pratense, respectively. Although sizes and growth stages ofplants and the composition of strigolactones produced weredifferent and thus direct comparisons were not possible, thetotal amount (or activity) of all strigolactones exuded byL. albus would be c. 1/1000 that exuded by T. pratense. Forexample, in the case of T. pratense, approx. 80% germinationof O. minor was induced at 500-µl equivalent of the controlculture (1/2 Tadano & Tanaka) medium (Yoneyama et al.,2007a) and a similar level of germination was achieved at450-ml equivalent of the control medium of L. albus (Fig. 4b).

Another clear difference between mycotrophic and nonmy-cotrophic plants was observed in the response of strigolactoneexudation to nutrient deficiency. In the cases of T. pratenseand S. bicolor, which are host plants of AM fungi, P deficiency(and also N deficiency in S. bicolor) significantly promotedstrigolactone exudation (Yoneyama et al., 2007a,b). In thecase of L. albus, however, P and N deficiency slightly reducedstrigolactone exudation (Fig. 4). Such a decrease in strigolactone

production may be attributable to a reduction in metabolicfunctions in L. albus under N and P deficiencies. In anothernonmycotrophic plant, Spinacia oleracia, neither P nor Ndeficiency enhanced strigolactone production (K. Yoneyama,unpublished).

Recently, the strigolactone orobanchol was identified fromthe root exudates of Arabidopsis thaliana, a host of Orobanchespp. but not of AM fungi (Goldwasser et al., 2008). Theseresults suggest that strigolactones have other unknown functionsindispensable for the normal growth and development ofplants themselves.

Legumes are thought to be very versatile in their symbioses(Sprent & James, 2007). Therefore, further work on thecharacterization of strigolactones from nodulating andnonnodulating legumes and legumes that produce ectomyc-orrhiza, and the effects of mineral nutrients on their strigolactoneproduction, can be expected to unveil the functions of strigo-lactones in the rhizosphere community and in plants.

Acknowledgements

Part of this study was supported by a Sasakawa ScientificResearch Grant from The Japan Science Society and a Grant-in-Aid for Scientific Research (1820810) from the Japan Societyfor the Promotion of Science (JSPS).

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