Strigolactone involvement in root development, response to ... · Strigolactone involvement in root development, response to abiotic stress and interactions with the biotic soil environment

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Strigolactone involvement in root development, response to abiotic stress and 1

interactions with the biotic soil environment 2

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Yoram Kapulnik and Hinanit Koltai 4

Institute of Plant Sciences, ARO, Volcani Center 5

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Corresponding Author: Yoram Kapulnik, [email protected]; Tel.: +972-50-6220461; Fax: 9

+972-3-9604180 10

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Running title: Strigolactone affect root development and response 15

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One Sentence Summary: Strigolactones, new plant hormones, play a role in root 18

development, root response to nutrient deficiency and plant interactions in the rhizosphere. 19

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Plant Physiology Preview. Published on July 18, 2014, as DOI:10.1104/pp.114.244939

Copyright 2014 by the American Society of Plant Biologists

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ABSTRACT 21

Strigolactones, recently discovered as plant hormones, regulate the development of different 22

plant parts. In the root, they regulate root architecture and affect root-hair length and density. 23

Their biosynthesis and exudation increase under low phosphate levels and they are associated 24

with root responses to these conditions. Their signaling pathway in the plant includes protein 25

interactions and ubiquitin-dependent repressor degradation. In the root, they lead to changes in 26

actin architecture and dynamics, and in localization of the PIN auxin transporter in the plasma 27

membrane. Strigolactones are also involved with communication in the rhizosphere. They are 28

necessary for germination of parasitic plant seeds, they enhance hyphal branching of arbuscular 29

mycorrhizal fungi and they promote rhizobial symbiosis. The current review focuses on the role 30

played by strigolactones in root development, their response to nutrient deficiency and their 31

involvement with plant interactions in the rhizosphere. 32

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INTRODUCTION 34

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Strigolactones have been recently discovered as plant hormones (Gomez-Roldan et al., 36

2008; Umehara et al., 2008) that are produced by a wide variety of plant species (Xie et al., 37

2010; Yoneyama et al., 2013). Several different types of strigolactones can be produced by a 38

single plant species, and different varieties of the same plant species may produce mixtures of 39

different types and quantities of strigolactone molecules (Xie et al., 2010; Yoneyama et al., 40

2013). Strigolactones are also produced in primitive plants, including Embryophyta and Charales 41

(Delaux et al., 2012). In all cases, they are produced and exuded in small amounts (e.g., Sato et 42

al., 2003; Yoneyama et al., 2007a; 2007b). Strigolactones are produced primarily in roots, but 43

their biosynthesis is not limited to the root system and also occurs in other plant parts (reviewed 44

by Koltai and Beveridge, 2013). 45

Although strigolactone biosynthesis derives from the carotenoid-synthesis pathway (Booker 46

et al., 2004; Matusova et al., 2005), only some of the proteins that are crucial for biosynthesis 47

have been identified to date. In the tested higher plant species, three plastid-localized proteins 48

have been found to be involved in the first stages of strigolactone biosynthesis (Booker et al., 49

2004, Matusova et al., 2005). One is a carotenoid isomerase, DWARF27 (D27), characterized in 50

rice (Oryza sativa L.), Arabidopsis (Arabidopsis thaliana) and pea (Pisum sativum) (Lin et al., 51

2009, Waters et al., 2012a; Adler et al., 2012). It can convert all-trans-β-carotene into 9’-cis-β-52

carotene (Alder et al., 2012). The latter is then oxidatively tailored, cleaved and cyclized by two 53

double-bond-specific cleavage enzymes, carotenoid cleavage dioxygenase (CCD) 7 and 8 54

(Booker et al., 2004; Schwartz et al., 2004), resulting in the bioactive strigolactone precursor 55

carlactone (Alder et al., 2012). The conversion of carlactone to strigolactone has not been 56

characterized, but may include MAX1, a class-III cytochrome P450 monooxygenase (Booker et 57

al., 2005; Alder et al., 2012; Cardoso et al., 2014). The presence of CCD enzymes has been 58

demonstrated in several diverse higher plants (Delaux et al., 2012). Moss (Physcomitrella 59

patens) also contains homologs of these three genes and accordingly, can produce strigolactones 60

(Proust et al., 2011). However, only some of these genes are present in other basal plants and 61

algae (Delaux et al., 2012). Approximately 15 strigolactones have been structurally characterized 62

to date (Ruyter-Spira et al., 2013); all consist of an ABC-ring system connected via an enol ether 63

bridge to a butenolide D ring (Xie et al., 2010). 64



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As plant hormones, strigolactones regulate the development of different plant parts. The 65

first indication that strigolactones function as plant hormones came from an examination of 66

hyperbranching mutants. These mutants' phenotype could not be attributed to altered levels of, or 67

response to one of the established plant hormones known at the time. Hence, a novel signal that 68

was associated with this phenotype was suggested (Beveridge et al., 1997). Later, this signal was 69

identified to be strigolactones, and to act as a long-distance branching factor that suppresses 70

growth of preformed axillary buds (Gomez-Roldan et al., 2008; Umehara et al., 2008). 71

Strigolactones dampen auxin transport in the main stem, thereby enhancing competition 72

between axillary branches and restraining axillary bud outgrowth (e.g., Bennett et al., 2006; 73

Mouchel and Leyser, 2007; Ongaro and Leyser, 2008; Crawford et al., 2010; Domagalska and 74

Leyser, 2011). Accordingly, strigolactones were demonstrated to act by increasing the rate of 75

removal of PIN1, the auxin export protein, from the plasma membrane of xylem parenchyma 76

cells in the stem. This activity was demonstrated by both computational model and experimental 77

data, and was correlated to the level of shoot branching observed in various mutant combinations 78

and strigolactone treatments (Shinohara et al., 2013). In pea, strigolactones were shown to induce 79

the expression of the bud-specific target gene BRANCHED1 (BRC1), which encodes a 80

transcription factor repressing bud outgrowth (Dun et al., 2012), and to be an auxin-promoted 81

secondary messenger (Dun et al., 2012; 2013; Brewer et al., 2009; Ferguson and Beveridge, 82

2009). Other activities of strigolactone include repression of adventitious-root formation 83

(Rasmussen et al., 2012) and plant height (de Saint Germain et al., 2013). They also induce 84

secondary growth in the stem (Agusti et al., 2011). Auxin positively regulates strigolactone 85

biosynthesis by elevating the expression of both MAX3 and MAX4. It has been suggested that 86

auxin and strigolactone modulate each other's levels and distribution, forming a dynamic 87

feedback loop between the two hormones (Hayward et al., 2009). 88

As noted, although the main site of strigolactone synthesis is the roots, part of their activity 89

is in the shoot. Therefore, strigolactones are expected be transported upward in the plant, from 90

root to shoot. Evidence to support this suggestion comes from Kohlen et al., (2011), who showed 91

the presence of the strigolactone orobanchol in the xylem sap of Arabidopsis. Another means of 92

strigolactone transport is probably via specific transporters. The Petunia hybrida ABC 93

transporter PDR1, localized mainly in the bud/leaf vasculature and subepidermal cells of the 94

root, was identified as a cellular strigolactone exporter. It was shown to regulate the level of 95



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symbiosis of arbuscular mycorrhizal fungi (AMF) (discussed further on) and axillary-shoot 96

branching (Kretzschmar et al., 2012). 97

Strigolactones also act in the root to determine root architecture. However, even before 98

strigolactones were identified as plant hormones, they were known to be involved with 99

communication in the rhizosphere. The current review focuses on strigolactone activity in the 100

roots as regulators of root-system architecture, root-hair length and primary root meristem, and 101

on aspects of their signaling. Their involvement with the root response to nutrient growth 102

conditions will also be presented and discussed. Moreover, the effects of strigolactone on root–103

rhizosphere communication will be presented, along with some implications on the evolution of 104

these interactions and their implementation. 105

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STRIGOLACTONES REGULATE ROOT DEVELOPMENT 107

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One of the first pieces of evidence suggesting that strigolactones have a role in the 109

development of root-system architecture was the finding that Arabidopsis mutants in the 110

strigolactone response or biosynthesis have more lateral roots than the wild type (WT; Kapulnik 111

et al., 2011a; Ruyter-Spira et al., 2011). Accordingly, treatment of seedlings with GR24 (a 112

synthetic and biologically active strigolactone; Johnson et al., 1976; Umehara et al., 2008; 113

Gomez-Roldan et al., 2008) repressed lateral root formation in the WT and strigolactone-114

synthesis mutants (max3 and max4), but not in the strigolactone-response mutant (max2), 115

suggesting that the negative effect of strigolactones on lateral root formation is MAX2-116

dependent (Kapulnik et al., 2011a; Ruyter-Spira et al., 2011). This negative effect on lateral root 117

formation was reversed in Arabidopsis under phosphate deficiency (Ruyter-Spira et al., 2011; 118

discussed further on). 119

Strigolactones are also suggested to regulate primary root length. GR24 led to elongation of 120

the primary root and to an increase in meristem cell number in a MAX2-dependent manner 121

(Ruyter-Spira et al., 2011; Koren et al., 2013). Accordingly, under conditions of carbohydrate 122

limitation, a shorter primary root and less primary meristem cells were detected in strigolactone-123

deficient and response mutants in comparison to the WT (Ruyter-Spira et al., 2011). 124

Furthermore, in rice, a major quantitative trait locus on chromosome 1—qSLB1.1—was 125

identified for the exudation of strigolactones, tillering, and induction of Striga germination 126



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(Cardoso et al., 2014). Several root-architectural traits were mapped in the same region (Topp et 127

al., 2013), suggesting that this locus may be involved in both strigolactone synthesis and root-128

system architecture. 129

Notably, expression of MAX2 under the SCARECROW (SCR) promoter was sufficient to 130

confer a response to GR24 in a max2-1 mutant background for both lateral root formation and 131

cell number in the primary root meristem (Koren et al., 2013). Since SCR is expressed mainly in 132

the root endodermis and quiescence center (Sabatini et al., 2003), these results point to an 133

important role for the endodermis in strigolactone regulation of root architecture. 134

Another one of strigolactones' effects in roots is on root-hair length. Exogenous 135

supplementation of various synthetic strigolactone analogs induced root-hair elongation in 136

Arabidopsis, in both the WT and strigolactone-deficient mutants (max3 and max4), but not in the 137

strigolactone-response mutant max2, suggesting that the effect of strigolactones on root-hair 138

elongation is mediated via MAX2 (Kapulnik et al., 2011a; Cohen et al., 2013). Furthermore, 139

response to auxin and ethylene signaling is required, at least in part, for the positive effect of 140

strigolactone on root-hair elongation. However, MAX2-dependent strigolactone signaling is not 141

necessary for the root-hair elongation induced by auxin (Kapulnik et al., 2011b). Hence, 142

strigolactones affect root-hair length at least in part through the auxin and ethylene pathways 143

(Koltai, 2011). Here too, expression of SCR::MAX2 was sufficient to confer root-hair elongation 144

in roots in response to GR24 (Koren et al., 2013). Since root-hair elongation is regulated in the 145

epidermis, the sufficiency of MAX2 expression under SCR (expressed mainly in the root 146

endodermis and quiescence center) for GR24 sensitivity suggests that strigolactones act non-cell-147

autonomously at short-range. 148

To summarize, strigolactones play a regulatory role in root development. At least part of this 149

activity is performed non-cell-autonomously, and may involve modulation of auxin transport, as 150

discussed further on. 151

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STRIGOLACTONE SIGNALING PATHWAY 153

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As indicated earlier for shoots, in the root, evidence also indicates a role for strigolactones in 155

the regulation of PIN protein activity. One piece of evidence comes from studies of tomato roots, 156

in which exogenous supplementation of 2,4-D (a synthetic auxin that is not secreted by auxin-157



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efflux carriers) led to reversion of the GR24-related root effect, suggesting functional 158

involvement of GR24 with auxin export (Koltai et al. 2010). Another piece of evidence comes 159

from studies in Arabidopsis, where treatment of seedlings with GR24 led to a decrease in PIN-160

FORMED (PIN1)–GFP intensity in lateral root primordia, suggesting that GR24 regulates PIN1 161

and modulates auxin flux in roots, and as a result, alters the auxin optima necessary for lateral 162

root formation (Ruyter-Spira et al., 2011). Furthermore, in Arabidopsis, following GR24 163

treatment that leads to root-hair elongation, PIN2 polarization was changed in the plasma 164

membrane of the root epidermis in the WT but not in the max2 mutant. In addition, in a MAX2-165

dependent manner, GR24 treatment led to increased PIN2 endocytosis, increased endosomal 166

movement in the epidermal cells, and changes in actin filament architecture and dynamics 167

(Pandya-Kumar et al., 2014). Together, these results suggest that strigolactones affect plasma 168

membrane localization of PIN proteins. At least for PIN2 in the root, they probably do so by 169

regulating the architecture and dynamics of actin filaments and PIN endocytosis, which are 170

important for PIN2 polarization (Pandya-Kumar et al., 2014; Figure 1). 171

Upstream of those events are probably those associated with strigolactone reception. One 172

of the components of strigolactone reception was identified several years ago as an F-box 173

protein, MAX2/D3/RMS4 (Stirnberg et al., 2002; Ishikawa et al., 2005; Johnson et al., 2006). An 174

additional component of strigolactone signaling is D14, which is a protein of the α/β-fold 175

hydrolase superfamily (Arite et al., 2009). Petunia DAD2, a homolog of D14, was shown to 176

interact in a yeast two-hybrid assay with petunia MAX2A only in the presence of GR24, 177

resulting in hydrolysis of GR24 by DAD2 (Hamiaux et al., 2012). In addition, in rice, D14 was 178

shown to bind to GR24 (Kagiyama et al., 2013) and to cleave strigolactones (Nakamura et al., 179

2013). 180

Moreover, via a Skp, Cullin, F-box (SCF)-containing complex (Moon et al., 2004), and in a 181

D14- and D3-dependent manner, it was shown in rice that strigolactones induce degradation of 182

D53, a class I Clp ATPase protein. D53 acts as a repressor of axillary bud outgrowth, and its 183

degradation by strigolactones prevents its activity in promoting axillary bud outgrowth (Jiang et 184

al., 2013; Zhou et al., 2013; Figure 1). Furthermore, in Arabidopsis, strigolactones were 185

suggested to induce, in a MAX2-dependent manner, proteasome-mediated degradation of D14 186

(Chevalier et al., 2014), suggesting a negative regulatory circuit of strigolactones and their own 187

signaling. 188



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This regulatory module of strigolactone/D14-like/D3-like/SCF is likely to have been 189

conserved in plant evolution (Waldie et al., 2014). As indicated above, it was shown to be 190

associated with strigolactone-regulated shoot development (Jiang et al., 2013; Zhou et al., 2013). 191

However it is not clear whether this or a similar reception system acts in the roots. It might be 192

that diversity in this module confers tissue specificity. Different D14-like proteins attached to 193

D3/MAX2 may confer different substrate specificity and as a result, a specific effect on plant 194

development. For example, a KAI2 (D14-LIKE)–MAX2-dependent pathway is responsible for 195

regulating seed germination, seedling growth and leaf and rosette development in response to 196

karrikins—strigolactone-analogous compounds originally found in forest-fire smoke (Flematti et 197

al., 2004; Waters et al., 2012b; Nelson et al., 2011; Waters et al., 2014). Modules for 198

strigolactone response that are composed of other α/β-fold hydrolases and/or degradation of 199

other repressors could potentially lead to execution of the strigolactone-related processes in roots 200

(Figure 1). 201

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STRIGOLACTONES ARE INVOLVED IN ROOT RESPONSES TO ABIOTIC STRESS 203

CONDITIONS 204

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Strigolactones seem to have been involved in plant responses to environmental stimuli 206

from their early evolution. In the moss P. patens, they determine the patterns of growth and 207

responses between neighboring colonies (Proust et al., 2011). In higher plants, they are involved 208

in both shoot and root architecture in response to nutritional conditions. 209

Pi is the inorganic form of phosphorus (P) that is available to plants. It is an essential 210

macronutrient for growth and development and in many places, it is considered to be a limiting 211

factor for growth (Bieleski, 1973; Maathuis, 2009). To cope with Pi deprivation, plants modify 212

their growth pattern and architecture. The shoot-to-root ratio is reduced under these conditions 213

(e.g., Ericsson, 1995); shoot branching is inhibited (reviewed by Domagalska and Leyser, 2011), 214

and root architecture is altered (Osmont et al., 2007; López-Bucio et al., 2003). Elongation of the 215

primary root is inhibited under conditions of Pi deficiency (Sánchez-Calderón et al., 2005) and 216

lateral root development is promoted (Nacry et al., 2005), probably for increased foraging of 217

subsurface soil. Following extended deprivation, root growth is also inhibited (Nacry et al., 218

2005). It should be noted, however, that these general patterns are not identical in all plant 219



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species. For example, under Pi deprivation, primary root growth is inhibited in some Arabidopsis 220

ecotypes but not in others (Chevalier et al., 2003). 221

Several plant hormones are known to regulate root-system architecture in response to 222

nutrient conditions. For example, under low Pi conditions, the changes in lateral root formation 223

in Arabidopsis have been suggested to result from increased auxin sensitivity, mediated by an 224

increase in the expression of the auxin receptor TIR1 (Pérez-Torres et al., 2008). Strigolactones 225

might be another plant hormone involved in the regulation of root-system architecture in 226

response to nutrient conditions. Although under conditions of sufficient Pi, strigolactones 227

negatively regulate lateral root formation (Kapulnik et al., 2011a), they reverse their effect to 228

positive regulation when Pi is limited (Ruyter-Spira et al., 2011). This suggests that 229

strigolactones act as another key regulator of lateral root formation, promoting their development 230

under low Pi conditions and repressing their emergence once Pi is abundant. 231

The length and density of root hairs are increased under Pi-deficient conditions, probably 232

to expand root surface area and enhance nutrient acquisition (Bates and Lynch, 2000; Péret et al., 233

2011; Gilroy and Jones, 2000). Indeed, the plant’s ability to absorb nutrients from the soil is 234

suggested to be directly associated with root-hair length and number (Sanchez-Calderon et al., 235

2005; reviewed by Gilroy and Jones, 2000). The recorded ability of strigolactone analogs to 236

increase root-hair length (Kapulnik et al., 2011a) may indicate their role in root-hair elongation 237

as an adaptive process in plants to growth conditions. 238

Also of significance is the dependence on strigolactones for the seedling response to Pi 239

deprivation, in terms of increasing root-hair density. Arabidopsis mutants, defective in 240

strigolactone biosynthesis or response, have a reduced ability to increase their root-hair density 241

in response to low Pi shortly after germination (Mayzlish-Gati et al., 2012). In accordance with 242

the suggestion that low Pi response is mediated by an increase in TIR1 expression (Pérez-Torres 243

et al., 2008), the strigolactone-response mutant, under conditions of Pi deprivation, displayed a 244

reduction, rather than induction of TIR1 expression (Mayzlish-Gati et al., 2012). 245

The reduced ability of strigolactone mutants to respond to low Pi conditions shortly after 246

germination may compromise survival of these seedlings under these conditions (Mayzlish-Gati 247

et al., 2012). These findings suggest an important role for strigolactones in plant adaptation to 248

stress. However, later on in plant development, even the strigolactone mutants recover, and are 249

able to respond to low Pi conditions (Mayzlish-Gati et al., 2012). This seedling recovery 250



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suggests the involvement of other mechanisms that are not dependent on strigolactones for 251

responding to Pi deprivation, which are effective later on in plant development. 252

Similarly, strigolactone involvement in responses to phosphate and nitrate was shown in 253

rice, by analyzing the response of strigolactone-synthesis (d10 and d27) or insensitive (d3) 254

mutants to reduced concentrations of Pi or nitrate (NO3−). Reduced Pi or NO3− concentrations 255

led to increased seminal root length and decreased lateral root density in the WT, but not in the 256

strigolactone mutants. Application of GR24 restored seminal root length and lateral root density 257

in the WT and in the strigolactone-biosynthesis mutants, but not in the strigolactone-response 258

mutant, suggesting that strigolactones are involved with the response to Pi and NO3− in rice as 259

well, leading to a D3-dependent change in rice root growth. In addition, based on changes in the 260

transport of radiolabeled indole-3-acetic acid, it was suggested that the mechanisms underlying 261

this regulatory role of D3/strigolactones involves modulation of auxin transport from shoots to 262

roots (Sun et al., 2014). 263

Pi deprivation leads to an increase in strigolactone exudation. Nitrogen (N) deficiency has 264

also been shown to increase strigolactone exudation. Nevertheless, it might be that N deficiency 265

affects strigolactone levels via its effect on P levels in the shoot. Indeed, a correlation was found 266

between shoot Pi levels and strigolactone exudation across plant species (Yoneyama et al., 267

2007a; 2007b; 2012). A clear correlation was also found in both Arabidopsis and rice between 268

this elevation in strigolactone levels and a decrease in shoot branching under restricted-Pi growth 269

conditions. In Arabidopsis, in correlation with the changes in shoot architecture, the level of the 270

strigolactone orobanchol in the xylem sap was increased under Pi deficiency (Kohlen et al., 271

2011). In rice, under these conditions, tiller bud outgrowth was inhibited and root strigolactone 272

(2′-epi-5-deoxystrigol) levels increased (Umehara et al., 2008). The increase in strigolactone 273

biosynthesis and exudation under low Pi conditions may also induce increased branching of 274

mycorrhizal hyphae (Akiyama et al., 2005; Besserer et al., 2006; 2008; Gomez-Roldan et al., 275

2008; Yoneyama et al., 2008), as detailed in the following chapters, and hence increased 276

mycorrhization. 277

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STRIGOLACTONES ARE SIGNALS FOR PLANT INTERACTIONS 282

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Strigolactones were initially identified as germination stimulants of the parasitic plants 284

Striga and Orobanche (e.g., Cook et al., 1972; Yokota et al., 1998; Matusova et al., 2005; Xie et 285

al., 2007; 2008a; 2008b; 2009; Goldwasser et al., 2008; Gomez-Roldan et al., 2008). It was only 286

later that strigolactones were also identified as stimulants of hyphal branching in AMF (e.g., 287

Akiyama et al., 2005; Besserer et al., 2006; 2008; Gomez-Roldan et al., 2008; Yoneyama et al., 288

2008). In addition, strigolactones were shown to stimulate nodulation in the legume–rhizobium 289

interaction process (Foo and Davies, 2011). 290

291

Mycorrhizal Symbiosis 292

The most prevalent symbiosis on earth is the arbuscular mycorrhizal (AM) symbiosis, which 293

consists of an association between the roots of higher plants and soil AMF. The AMF are 294

members of the fungal phylum Glomeromycota (Redecker and Raab, 2006), and symbiotic 295

associations are formed with most terrestrial vascular flowering plants (Smith and Read, 2008), 296

in most cases contributing to plant development, especially under suboptimal growth conditions. 297

During the symbiosis, AMF hyphae, which extend through the soil, provide a greater root 298

surface area to exploit a larger volume of soil, thereby enhancing the amount of nutrients 299

absorbed from the soil for the plant (Rausch and Bucher, 2002). In return, the fungus receives 300

fixed carbon in the form of glucose (reviewed by Douds et al., 2000), hexoses (Shachar-Hill et 301

al., 1995; Solaimand and Saito, 1997) or sucrose from the host. 302

In general, the AM symbiosis is comprised of two distinct functional stages: the 303

"presymbiotic stage" and the "symbiotic stage". A very detailed description of the presymbiotic 304

stage confirmed that fungal spore germination in the soil and the growth of fungal hyphae are 305

both stimulated in the presence of a host root (Mosse and Hepper, 1975; Gianinazzi-Pearson et 306

al., 1989; Giovannetti et al., 1996; Buée et al., 2000; Nagahashi and Douds, 2000; Requena et al., 307

2007). These two phenomena, together with the hyphal branching response, may reflect unique 308

communication in the rhizosphere to enhance successful mycorrhization on the host (Koske and 309

Gemma, 1992). 310

Purified strigolactones from root exudates or synthetic strigolactones have been shown 311

capable of inducing hyphal branching in many AMF (Akiyama et al., 2005). These molecules are 312



12

present at sub-nanogram levels in the rhizosphere (Akiyama and Hayashi, 2006; Akiyama et al., 313

2005). Similarly, the synthetic strigolactone GR24 was shown to effectively induce AMF hyphal 314

branching at a concentration of 10-8 M (Gomez-Roldan et al., 2008). Accordingly, strigolactone-315

deficient mutants of pea and tomato exhibit reduced levels of AMF hyphal branching in their 316

rhizosphere as compared to the response obtained in the presence of WT root exudates (Gomez-317

Roldan et al., 2008; Koltai et al., 2010b). 318

In a more in-depth study, it was demonstrated that strigolactones rapidly induce changes in 319

AMF energy metabolism before any gene-expression process in the fungus can be detected. 320

When AMF hyphae were exposed to the synthetic strigolactone GR24, a rapid alteration (within 321

60 min) in mitochondrial shape, density and motility was observed. In Gigaspora rosea hyphae, 322

NADH concentrations, dehydrogenase activity and ATP content were altered within minutes by 323

application of GR24 (Besserer et al., 2006; 2008). 324

The importance of strigolactones to AMF establishment was reinforced by the observation 325

of a lower colonization rate of a tomato strigolactone-biosynthesis mutant by AMF spores than 326

that obtained in WT roots. Interestingly, these differences were less pronounced when plants 327

were inoculated with "whole inoculum" (consisting of spores, hyphae and infected roots) (Koltai 328

et al., 2010b). 329

Nevertheless, it was shown that root exudates of mycorrhitic plants induce Striga and 330

Orobanche seed germination to a lesser extent than the induction obtained by exudates of non-331

mycorrhitic plants (Lendzemo et al., 2009; Fernández-Aparicio et al., 2010). Moreover, 332

strigolactone production was shown to be significantly reduced in roots of mycorrhitic tomato 333

plants (López-Ráez et al., 2011). Therefore, strigolactones may be negatively regulated by AMF 334

via a feedback loop. Alternatively, it may be that AMF colonization has a significant impact on 335

enhancement of AM symbiosis and consequently, increased Pi acquisition, which may then be 336

reflected in elevated levels of P content, the latter inducing suppression of strigolactone 337

biosynthesis. 338

It is not yet clear whether strigolactones have any role during fungal morphogenesis in the 339

host cortical cells or during symbiosis stages. Moreover, it is not clear whether strigolactones are 340

essential for the AMF interaction. 341

342

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Parasitic Plants 344

Witchweed (Striga spp.) and broomrape (Orobanche and Phelipanche spp.) are important 345

parasitic weeds that have a devastating effect on the production of many crop species, resulting 346

in economic damage and food losses worldwide (Parker, 2009). The damage conferred on crop 347

development and productivity has been summarized elsewhere (Joel, 2000; Gressel and Joel, 348

2013). 349

The communication between parasites and their host plants depends on strigolactones, as 350

signal molecules which are exuded from the host roots into the rhizosphere. These signals mostly 351

involve induction of seed germination of the parasitic plants which, within a few days, must 352

attach to their host to acquire nutrients, or die (Joel and Bar, 2013). This communication allows 353

germination of the parasitic plant at the right distance from the root surface, at the right time in 354

the season and with the right nutritional, temperature and moisture levels in the host rhizosphere. 355

Following the parasitic plant's germination, a tubercle develops underground and shoot 356

outgrowth is initiated. The shoots emerge aboveground, flower, and produce tens of thousands of 357

seeds (for a more in-depth review, the reader is referred to Joel, 2013). 358

Three types of isoprenoid compounds stimulate the germination of root-parasitic plants: 359

dihydrosorgoleone, the sesquiterpene lactones, and strigolactones (Bouwmeester et al., 2003). 360

Strigolactones are active at extremely low concentrations (on the order of 10-7 to 10-15 M; Joel, 361

2000). A variety of natural strigolactones were shown able to induce germination of Orobanche 362

minor from 10 pM strigolactones (for orobanchol, 2'-epiorobanchol and sorgomol) to 10 nM for 363

7-oxoorobanchol. The synthetic analog GR24 is 100-fold less active than natural strigolactones 364

(Kim et al., 2010). Advances in chromatography and mass spectrometry are enabling the 365

discovery and characterization of novel strigolactones. 366

367

Symbiotic Interactions with Rhizobium 368

The nitrogen-fixing bacteria of the genus Rhizobium play a fundamental role in nodule 369

formation and beneficial symbiotic interactions on the roots of legumes such as pea, bean 370

(Phaseolus vulgaris), clover (Trifolium spp.) and alfalfa (Medicago sativa). The symbiotic 371

interaction involves signal exchange between the partners that leads to mutual recognition and 372

development of the nodule structure. In short, at the preinfection stage, the bacteria sense 373

flavonoids that are secreted from the legume host roots. These flavonoid molecules, which are 374



14

specific to each legume–rhizobium interaction, activate the production and secretion of 375

lipochitooligosaccharide nodulation (Nod) factors from the bacteria that are recognized by the 376

host plant. In most cases, Nod-factor perception leads to induction of a cascade of signaling 377

events, such as root-hair deformation and infection-thread formation, and is involved in triggering 378

cell division in the cortex of the root, leading to nodule organ formation. Production of Nod 379

factors and exopolysaccharides by many of the rhizobium bacteria elicits an infection thread 380

where the penetrated bacteria multiply. Intracellular bacterial cells then differentiate to 381

"bacteroids"—the nitrogen-fixing stage of the symbiosis. In addition to the morphological and 382

cellular modifications, plants and bacteria produce and respond to large groups of peptides, 383

transcriptional factors, and early and late nodulation (nod) genes. However, the molecular 384

mechanisms by which many of them are involved in the differentiation process are poorly 385

understood. Within the nodule meristem and after cell differentiation, rhizobia bacteroids supply 386

ammonia or amino acids to the plant and in return, receive organic acids as a carbon and energy 387

source (Markmann and Parniske, 2009). 388

The ability of strigolactones to alter plant-meristem development led to the search for 389

strigolactone involvement in this symbiotic interaction. The potential of strigolactones to control 390

nodulation was verified using the strigolactone-deficient rms1 mutant in pea (Pisum sativum L.) 391

(Foo and Davis, 2011). This work showed that endogenous strigolactones are positive regulators 392

of nodulation in this plant. Using rms1 mutant root exudates and root tissue that were almost 393

completely deficient in strigolactones, Foo and Davies, (2011) demonstrated a 40% reduction in 394

the number of nodules relative to WT plants that contained strigolactones. Application of GR24 395

to rms1 plants resulted in an increase in the number of nodules on the mutant roots to the level 396

obtained on the WT (without exogenous application of strigolactones). GR24 application can also 397

enhance nodule number in WT pea, M. sativa and Lotus japonicus (Soto et al., 2010; Foo and 398

Davies, 2011; Liu et al., 2013). It was also shown that strigolactones in the root, but not shoot-399

derived factors, can regulate nodule number (Foo et al., 2014). Genetic studies indicated that 400

strigolactones may act relatively early in nodule formation, rather than during nodule 401

organogenesis (Foo et al., 2013 a, b). Moreover, it was shown that strigolactones do not influence 402

nodulation by acting directly on the rhizobium bacteria (Soto et al., 2010) or on the Ca2+ 403

signaling that follows flavonoid perception (Moscatiello et al., 2010). It was suggested that 404

strigolactones are not essential for the development of a functional nodule but may be important 405



15

in determining the optimal nodule number, thereby having a quantitative effect on nodulation in 406

pea (Foo et al., 2014). 407

The mechanism governing strigolactone enhancement of the nodulation process is still an 408

enigma. One potential explanation might be that strigolactones interact with auxin distribution 409

during the cell-division process that leads to nodule development. Auxin was found to be 410

accumulated in dividing cortical cells in L. japonicus and NODULE INCEPTION, a key 411

transcription factor in nodule development, was found to positively regulate this accumulation 412

(Suzaki et al., 2012). Thus, the fact that strigolactones also regulate auxin transport suggests an 413

additional regulatory path for this symbiotic interaction. Furthermore, it was shown that auxin 414

positively regulates strigolactone production (Hayward et al., 2009), which suggests a positive-415

feedback regulation loop between auxin and strigolactones in the promotion of nodulation in 416

legume roots. 417

418

CONCLUDING REMARKS 419

420

Strigolactones are an important group of molecules. They likely developed in plants as key 421

regulators of plants' developmental adaptations to environmental conditions. Their production 422

and exudation from roots was utilized by other organisms to the benefit (AMF and rhizobia) or 423

detriment (parasitic plants) of the host plants. Strigolactones may be involved in additional cases 424

of communication in the rhizosphere, as well as in additional responses of the plant to growth 425

conditions. This is due to the high complexity of the rhizosphere (Jones and Hinsinger, 2008), 426

which is composed of (1) a multiplicity of organisms and (2) microconditions in soil “pockets”, 427

e.g., low levels of nutrients. This complexity may hinder the evaluation of additional functions of 428

strigolactones under the non-homogeneous rhizospheric conditions that are normally found in 429

nature. 430

Another interesting aspect of strigolactones is their potential use in agriculture. A number of 431

publications have discussed their implementation as inducer of suicidal seed germination of 432

parasitic plant (e.g., Zwanenburg et al., 2009; Vurro and Yoneyama, 2012; Zwanenburg et al., 433

2013). However, strigolactones may be used in additional approaches for the development of 434

new agricultural methodologies and technologies, compatible with emerging concepts of 435

sustainable agriculture. For example, strigolactones may be used for improvement of root-system 436



16

architecture, e.g., to develop a hyperbranched root system for increased nutrient-use efficiency, 437

or deeper roots for increased water-use efficiency. They may also be used for regulation of shoot 438

branching when, for example, the emergence of axillary branches is undesirable. Strigolactone 439

analogs and mimics that are specific for one activity (e.g., shoot branching) are already being 440

developed (Fukui et al., 2011; 2013; Boyer et al., 2013). Strigolactone inhibitors may be used to 441

enhance rooting of plant cuttings (Rasmussen et al., 2012), potentially promoting the propagation 442

of woody plants for the industry and for conservation of endangered species. Today's new 443

strigolactone analogs and mimics, which are under development or being synthesized 444

(Zwanenburg et al., 2009; Prandi et al., 2011), are likely to substantially promote the ability to 445

use strigolactones to the benefit of agriculture. 446

447

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813

814

815

816

817

Figure legend 818

819

Figure 1. A model of the putative signaling pathway of strigolactone in roots. An α/β-fold 820

hydrolase protein may serve as the strigolactone receptor. It may interact with MAX2 and as a 821

result, lead to degradation by ubiquitination of a repressor (reviewed by Waldie et al., 2014). 822

These or similar events may lead to changes in the architecture and dynamics of actin filaments 823

and PIN endocytosis which is important for PIN2 polarization. As a result, PIN2 protein 824

polarization is affected, which may lead to changes in auxin flux, and to execution of 825

strigolactone-associated root effects, such as root-hair elongation. 826

827

828



MAX2 SCF complex

α/β HYDROLASE REPRESSOR

Root responses (e.g., root hair elongation)

Strigolactones

Ubiquitin

PIN containing membrane

Actin filament

?

Key



Strigolactone involvement in root development, response to ... · Strigolactone involvement in root development, response to abiotic stress and interactions with the biotic soil environment

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