Towards secure 5G networks: A Survey

Received: 14 March 2019

Revised: 27 July 2019

Accepted: 30 July 2019

DOI: 10.1111/pce.13636

ORIGINAL ARTICLE

Flowering‐mediated root‐fungus symbiosis loss is related to jasmonate‐dependent root soluble sugar deprivation Wei Zhang

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Jie Yuan

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Ting Cheng

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Meng‐Jun Tang

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Kai Sun

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Shi‐Li Song

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Fang‐Ji Xu

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Chuan‐Chao Dai Jiangsu Key Laboratory for Microbes and Functional Genomics, Jiangsu Engineering and Technology Research Center for Industrialization of Microbial Resources, College of Life Sciences, Nanjing Normal University, Nanjing, China Correspondence Chuan‐Chao Dai; Jiangsu Key Laboratory for Microbes and Functional Genomics, Jiangsu Engineering and Technology Research Center for Industrialization of Microbial Resources, College of Life Sciences, Nanjing Normal University, Nanjing, China. Email: [email protected] Funding information National Natural Science Foundation of China, Grant/Award Number: NSFC 31870478; Priority Academic Program Development (PAPD) of Jiangsu Higher Education Institutions of China; Doctor Breeding Project of Nanjing Normal University, Grant/Award Number: 1812000006317

Abstract The role of flowering in root‐fungal symbiosis is not well understood. Because flowering and fungal symbionts are supported by carbohydrates, we hypothesized that flowering modulates root‐beneficial fungal associations through alterations in carbohydrate metabolism and transport. We monitored fungal colonization and soluble sugars in the roots of Arabidopsis thaliana following inoculation with a mutualistic fungus Phomopsis liquidambari across different plant developmental stages. Jasmonate signalling of wild‐type plants, sugar transport, and root invertase of wild‐type and jasmonate‐insensitive plants were exploited to assess whether and how jasmonate‐dependent sugar dynamics are involved in flowering‐mediated fungal colonization alterations. We found that flowering restricts root‐fungal colonization and activates root jasmonate signalling upon fungal inoculation. Jasmonates reduce the constitutive and fungus‐induced accumulation of root glucose and fructose at the flowering stage. Further experiments with sugar transport and metabolism mutant lines revealed that root glucose and fructose positively influence fungal colonization. Diurnal, jasmonate‐dependent inhibitions of sugar transport and soluble invertase activity were identified as likely mechanisms for flowering‐mediated root sugar depletion upon fungal inoculation. Collectively, our results reveal that flowering drives root‐fungus cooperation loss, which is related to jasmonate‐dependent root soluble sugar depletion. Limiting the spread of root‐fungal colonization may direct more resources to flower development. K E Y W OR D S

Arabidopsis thaliana, flowering, jasmonate, Phomopsis liquidambari, sugar, symbiosis

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I N T RO D U CT I O N

leaves and roots are the major C sinks at vegetative stage, and flowers are new and strong C sinks during the transition from vegetative to

Plants constantly modulate their growth and development to react

reproductive growth (Borghi & Fernie, 2017). Flowers compete with

and adjust to changing environmental conditions. Partitioning of car-

roots and other heterotrophic tissues for C based on their relative C

bohydrates between source and sink organs is one of the major deter-

sink strength (Borghi & Fernie, 2017; Müller, Drincovich, Andreo, &

minants of plant growth and development (Durand et al., 2018; Koch,

Lara, 2010). Emerging evidences suggest that plants modulate

2004). Photosynthesis fixed carbon (C) sustains the growth and devel-

flowering time through changing endogenous C status to reduce the

opment of nonphotosynthetic sinks, including young leaves, stems,

risks imposed by a stressful environment (Kazan & Lyons, 2016; Kim,

flowers, roots, and seeds (Koch, 2004). For flowering plants, young

Kim, & Park, 2007; Lauxmann et al., 2016).

Plant Cell Environ. 2019;1–19.

wileyonlinelibrary.com/journal/pce

© 2019 John Wiley & Sons Ltd

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Plants are engaged with a wide range of beneficial fungi that pro-

Klempien, & Dudareva, 2014). Interference in sugar metabolism

vide plant with various benefits, for instance, enhanced root system

results in early arrest of floral development, thereby leading to prema-

growth (Sukumar et al., 2013; Zhang et al., 2018), increased water

ture flowering and sterility (Lauxmann et al., 2016; Seo, Ryu, Kang, &

and nutrient uptake (Almario et al., 2017; Ezawa & Saito, 2018), and

Park, 2011). For instance, INDETERMINATE DOMAIN 8 (idd8)‐

better stress tolerance (Sui, Zhang, Tian, Xue, & Li, 2019) in exchange

silenced plants exhibit early flowering because of compromising sugar

for photoassimilates. Root beneficial fungi have also been demon-

transport and metabolism (Seo et al., 2011). Because both fungal

strated to influence the timing of important developmental transitions,

symbionts and flowers are fed on carbohydrates, if plant flowering

especially flowering (Das et al., 2012;Pan et al., 2017; Zavala‐Gonzalez

modulates root fungal colonization, the whole plant's carbohydrate

et al., 2017). For example, the colonization by root endophytes

partitioning and metabolism may be involved in this process.

Piriformospora indica and Pochonia chlamydosporia stimulate the

Flowering has been demonstrated to modulate plant hormones to

expression of key flowering time regulators and shortens the

coordinate metabolic networks, of which jasmonates play an impor-

flowering time of Arabidopsis (Pan et al., 2017; Zavala‐Gonzalez

tant role (Diezel, Allmann, & Baldwin, 2011; Li et al., 2017; Stitz, Hartl,

et al., 2017). Despite cumulative laboratory and field experiments

Baldwin, & Gaquerel, 2014). Jasmonates have been implicated in floral

showing that fungal symbionts induce plant precocious flowering,

development and opening, and regulating the primary and secondary

the role of flowering on root‐beneficial fungal mutualisms has rarely

metabolism in open flowers (Li, Wang, et al., 2017; Stitz et al., 2014).

been investigated.

Jasmonates have been intensively studied as positive regulators of

The root accessible carbohydrate milieu influences the coloniza-

plant secondary metabolism in pathogen and herbivore resistance

tion degree of fungal symbionts (Schaarschmidt, Kopka, Ludwig‐

(Howe & Jander, 2008; Okada, Abe, & Arimura, 2015). In addition to

Müller, & Hause, 2007; Vargas, Crutcher, & Kenerley, 2011; Vargas,

secondary metabolism, jasmonates have also been implicated in regu-

Mandawe, & Kenerley, 2009). The level of accessible carbohydrate in

lating plant primary metabolism. Herbivore Manduca sexta attacks

root tissue is not constant and varies depending on plant species, plant

reduce leaf glucose and fructose levels of Nicotiana attenuata in a

developmental stage, photosynthetic activity, root traits, etc. Sugars

jasmonate‐dependent manner (Machado, Arce, Ferrieri, Baldwin, &

are generated in source leaves and transported to roots by the Sugars

Erb, 2015). JA‐methyl ester (MeJA) application reduces levels of starch

Will Eventually be Exported Transporter (SWEET) and Sucrose Uptake

and sugars in leaves and roots (Machado et al., 2013), indicating that

Transporter families (Chen et al., 2010, 2012; Doidy et al., 2012). After

jasmonates negatively influence primary metabolism. Moreover,

reaching the root tissue, plant‐derived invertase hydrolyses sucrose to

jasmonates have been found to be crucial role for the establishment

hexose, which feeds the fungal symbionts (Doidy et al., 2012). Studies

and

have demonstrated that the availability of hexose in apoplast controls

reconfiguring local and systemic metabolism (Gerlach et al., 2015;

the extent of plant–fungal associations (Schaarschmidt et al., 2007;

Hause & Schaarschmidt, 2009; Landgraf, Schaarschmidt, & Hause,

Vargas et al., 2009). In addition to being energy sources for root fungal

2012). Indeed, transcriptomic and metabolomic studies showed acti-

symbionts, sugars also act as signalling molecules for plants, and thus

vated JA signalling and metabolic rearrangement within roots and

plants strictly modulate transport and redistribution of sugars to

shoots of multiple plant species upon fungal inoculation (Adolfsson

ensure their growth, development, and reproduction (Ruan, 2014).

et al., 2017; Gerlach et al., 2015). Plants deficient in jasmonate biosyn-

Despite the reciprocal benefits, plant–beneficial fungal interactions

thesis or signalling display high level of fungal colonization (Zavala‐

may range from mutualism to parasitism based on the cost–benefit

Gonzalez et al., 2017). Although the exact roles of jasmonates in

trade‐offs. If the fungal growth within the root is out of control,

plant–fungal associations are still unclear, several studies have

root‐fungal mutualism could switch to parasitism or even pathogene-

reported that jasmonate signalling restricts fungal root spread through

sis, leading to stunted plant growth (Johnson, Graham, & Smith,

limiting the flow of carbohydrate to fungi (Landgraf et al., 2012;

1997; Kogel, Franken, & Hückelhoven, 2006; Zavala‐Gonzalez et al.,

Zavala‐Gonzalez et al., 2017). Given the importance of carbohydrate

2017). Therefore, plants limit the spread of fungal partners when

in plant flowering and root‐fungal symbioses, as well as of

encountering carbohydrate conflicts. For example, the colonization

jasmonate‐mediated carbohydrate changes in root‐fungal mutualisms,

of root‐associated symbionts was inhibited by long‐lasting herbivory

we hypothesized that plant flowering and root fungal colonization may

attacks that induced foliage loss by alterations in above‐ground and

be linked through jasmonate‐dependent carbohydrate changes.

maintenance

of

plant–beneficial

fungal

interactions

by

below‐ground allocation of carbohydrates (Gange, Bower, & Brown,

The root endophyte Phomopsis liquidambari, which is horizontally

2002; Markkola et al., 2004). Shaded Medicago sativa showed reduced

transmitted via the environment (Rodriguez, White, Arnold, &

mycorrhizal colonization because of limited carbohydrates supply to

Redman, 2009), establishes beneficial associations with morphologi-

the fungal partners (Fellbaum et al., 2014). During the plant's life cycle,

cally and biochemically different hosts (Zhang et al., 2016; Zhang

flowering is a critical developmental event that must be tightly regu-

et al., 2018; Zhou, Li, Chen, & Dai, 2017). Phomopsis liquidambari‐

lated to maximize seed bearing success (Kazan & Lyons, 2016).

treated crop plants showed an increase in nutrient uptake (Li et al.,

Flowers are energy intensive sinks that draw carbohydrates and nutri-

2017; Yang et al., 2014), resistance against pathogens (Xie et al.,

ents towards them (Borghi & Fernie, 2017; Lauxmann et al., 2016). Pri-

2017), and improved yield (Zhang et al., 2017). Moreover, rice infected

mary metabolites sustain pollen development, pollen tube growth,

with P. liquidambari underwent precocious flowering by 4 days and the

pollination, and nectar formation (Borghi & Fernie, 2017; Muhlemann,

whole growth period was completed approximately 10 days early (Li,

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Zhou, et al., 2017). Our previous study with P. liquidambari and

according to the method described by Camehl et al. (2010). The mock

Arabidopsis

treatment received same amount of heated‐killed mycelium.

thaliana

(hereafter

Arabidopsis)

showed

that

the

colonization rate of fungi in roots was gradually increased during the vegetative stage and then decreased at the reproductive stage (Zhang

2.2

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Quantification of fungal colonization

et al., 2018), suggesting a possible role of plant flowering in modulating fungal symbiosis. Additionally, in vitro and in vivo experiments

Arabidopsis plants were collected and open flowers (stages 13 and 14)

indicated that soluble sugar modulates colonization of P. liquidambari

were counted (Figure 3a; Smyth, Bowman, & Meyerowitz, 1990;

(Sun et al., 2019).

Widemann, Smirnova, Aubert, Miesch, & Heitz, 2016), and then

The aim of the present study is to explore whether and mecha-

divided the plants into shoots and roots. Flowers at stage 13 and 14

nisms by which plant flowering affect root‐fungal mutualism with

were chosen because they are fully heterotrophic organs (Borghi &

Arabidopsis–P. liquidambari system. First, we asked whether plant

Fernie, 2017; Thomas, Ougham, Wagstaff, & Stead, 2003). The extent

flowering regulates root P. liquidambari colonization. We then analyzed

of P. liquidambari root colonization of the plants was determined by

the jasmonates signalling and root soluble sugar to determine whether

the methods described by Peskan‐Berghöfer et al. (2015) with P.

jasmonate‐dependent sugar dynamics are involved in plant flowering‐

liquidambari specific ITS primer set (Bf1/Br1; Zhang et al., 2018) or

mediated fungal colonization alterations. Lastly, diurnal changes of

GFP with Arabidopsis AtUBQ5 (At3g62250; Khatabi et al., 2012) as a

root soluble sugar, transport of soluble sugar in phloem, and activity

reference gene.

of root invertases in wild‐type (WT) and jasmonate‐insensitive plants

The Arabidopsis root system was sampled at 12 and 20 days after

were exploited to reveal the likely mechanisms underlying flowering‐

inoculation (dai) and processed as described by Zhou et al. (2018) for

mediated root sugar depletion upon fungal inoculation.

confocal laser scanning microscopy (Nikon, Tokyo, Japan). Images were processed with Adobe Photoshop 7.0 (CA, USA).

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MATERIALS AND METHODS

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2.1

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Plant and fungal materials

Arabidopsis thaliana WT and all mutant lines were in the Columbia (Col‐0) background. The seeds of coi1‐2 (Zhu et al., 2011), myc2‐2 (Zhu et al., 2011), and 35Spro:MYC2 (Chen et al., 2011) were provided by Ziqiang Zhu, Nanjing Normal University. Homozygous A. thaliana insertion mutant lines flc (European Arabidopsis Stock Centre code N541126), lfy (European Arabidopsis Stock Centre code N6228), sweet11 (European Arabidopsis Stock Centre code N680842), sweet12 (European Arabidopsis Stock Centre code N685601), suc1 (European

2.3 | Effect of P. liquidambari on Arabidopsis growth, development, and yield After transplanting, the days to bolting, flowering and first silique shattering were recorded. The numbers of days from sowing to bolting to 1 cm of main stem, appearance of first floral opening, and first silique shattering were scored as time of bolting, flowering, and first silique shattering, respectively (Zavala‐Gonzalez et al., 2017). Meanwhile, rosette leaves at bolting, total leaves at flowering, inflorescences at 40 dai and silique number at 45 dai were also counted. At the finial harvest, the seeds were collected and the yields were recorded.

Arabidopsis Stock Centre code N698444), suc2 (European Arabidopsis Stock Centre code N698644), cinv1 (European Arabidopsis Stock

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Quantification of JA and JA‐Ile concentrations

Centre code N2104101), and cwinv2 (European Arabidopsis Stock Centre code N687578; Alonso et al., 2003) were obtained from

The extraction and detection of JA and JA‐Ile concentrations were as

European Arabidopsis Stock Centre. The seeds were surface sterilized

according to Mao et al. (2017). The JA and JA‐Ile in leaf and root were

and stratified at 4°C for 48 hr before they were sown on a mix of ster-

determined using an LC/MS system with a C18 column (Agilent

ilized vermiculite and peat (2:1; v/v) as described in Zavala‐Gonzalez

Technologies, USA) with 0.05 % formic acid (A) and methanol (B) as

et al. (2017). The seeds were germinated in a greenhouse under long

the mobile phase, and dihydro‐JA was used as internal standard

day (23°C, 16 hr light/8 hr dark, 60% relative humidity). Eight‐day‐

(Zavala‐Gonzalez et al., 2017).

old uniform seedlings were carefully transferred to 250‐ml pots with approximately 200 g of same substrate. The pots were randomly distributed and watered with sterilized distilled water as required.

2.5 | Quantification of soluble sugars, starch and protein concentrations in Arabidopsis roots

P. liquidambari strain B3 was isolated from the inner bark of Bischofia polycarpa (Chen, Xie, Ren, & Dai, 2013) and stored and acti-

To detect the concentrations of soluble sugars, starch, and protein,

vated according to Zhang et al. (2018). It was labelled with green fluo-

200‐mg roots were sampled at 14:00 hr and extracted with 80%

rescent protein (GFP) by the pCT74 plasmid and has been proven to

ethanol. After incubation at 80°C for 20 min, samples were centri-

act as a root endophyte of rice and peanut (Zhang et al., 2018; Zhou,

fuged at 12,000 g for 20 min and the supernatants were collected.

Li, Huang, & Dai, 2018). Before seedlings were transferred to pots, the

The pellets were reextracted twice with the same procedures, and all

growth substrate was mixed carefully with the mycelium (1% w/w)

supernatants were combined for sucrose, glucose, and fructose

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determinations. The remaining pellets were used for starch determina-

differences. Correlations were identified by Pearson correlation analy-

tion. The concentrations of soluble sugars and starch were detected

sis. Graphs and images were assembled with Adobe Photoshop 7.0

with assay kits (Nanjing Jiancheng Bio‐engineering Institute, Nanjing,

(CA, USA).

China; Zhang et al., 2017). The content of protein was quantified by the Bradford method (Bradford, 1976).

3 2.6 | Quantification of phloem sugar transport and root invertase activity To reveal the mechanisms underlying jasmonate‐dependent sugar dynamics upon P. liquidambari inoculation at anthesis, we analysed the root soluble sugars, phloem sugar transport, and root invertase activity. Because carbohydrate concentrations, phloem sugar transport, and invertase activity show diurnal variation patterns (Flis et al., 2019; Machado et al., 2015), roots were sampled at five time points during the day: 08:00, 11:00, 14:00, 18:00, and 22:00 hr to determine the soluble sugar concentrations in root tissue and phloem sap, and the activity of soluble and insoluble invertases in root tissue. The invertase enzyme activities were analysed by methods described by Ferrieri et al. (2015). Rosette leaves were collected at flowering time for phloem sap analysis, and their fresh weight was recorded. After cutting at the base of the petioles, the petioles were immersed in phosphate buffer (50 mM, pH 7.5) containing 10‐mM ethylenediaminetetraacetic acid (Vilaine et al., 2013). Phloem sap exudates were collected in the dark (60% RH) for 2 hr and used for sucrose, glucose, and fructose content determinations.

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RESULTS

3.1 | Phomopsis liquidambari colonizes Arabidopsis roots At 12 and 20 dai, abundant green fluorescent hyphae were found on primary root surfaces of Arabidopsis (Figure 1a and 1e). After close inspection at 20 dai, the hyphae maintained their macroscopic structure and texture and formed “runner hyphae” on the root surface (Figure 1b). Runner hyphae penetrated the epidermal cells through forming thicker bulbous invasive hyphae at the hyphopodia‐like infection structures (Figure 1c, 1d, 1f, and 1g). The outer cuticle of the plant was obviously squeezed or breached causing by hyphal constriction at the infection sites, but plant cells remained intact and root necrosis was not observed (Figure 1c, 1d, and 1f). Occasionally, we found branched hyphopodia‐like infection structures during the infection process (Figure 1g). In addition, runner hyphae also entered the root tissue through crack sites of surface (Figure 1h). After epidermis penetration, some hyphae continued to growth and reached the root inner cortex (Figure 1i). We also observed abundant green fluorescent hyphae on lateral roots (Figure 1j and 1k). Consistent with our previous study (Zhang et al., 2018), P. liquidambari was confined to root tissues, whereas shoots of Arabidopsis were free of hyphae (data

2.7 | Plant RNA extraction and quantitative real‐time‐polymerase chain reaction analysis For plant RNA extraction, approximately 150 mg rosette leaves and roots were sampled. The RNA extraction was performed with TRIzol reagent (Vazyme Biotech Co., Ltd.) according to the manufacturer's instructions. The RNA was treated with DNase I to remove genomic DNA. The synthesis of first‐strand cDNA was performed with a Reverse Transcription System Kit (Vazyme Biotech Co., Ltd.). Quantitative real‐time‐polymerase chain reaction (RT‐PCR) was carried out on a 7500 RT‐PCR System (Applied Biosystems) using AceQ qPCR SYBR® Green Master mix (Vazyme Biotech Co., Ltd.), according to the methods described by Zhang et al. (2018). AtACTIN2 (At3g18780) was used as reference gene. The experiment was conducted at least three independent replicates. The primers used in the present study are listed in Table S1 in the supporting information. The relative expression of target gene was determined by the △Ct method.

not shown). Overall, these results suggest that P. liquidambari acts as a root endophyte of Arabidopsis.

3.2 | Phomopsis liquidambari promotes growth, flowering, and yield of Arabidopsis To investigate the effects of P. liquidambari on growth and development of Arabidopsis, a long‐term pot experiment was performed. Plants colonized by P. liquidambari exhibited bigger leaves and higher chlorophyll a content than those in mock controls at 12 dai (Figure 2j, 1k, and S1 in the supporting information). No significant difference was observed in bolting time in response to P. liquidambari inoculation (Figure 2a). By contrast, P. liquidambari promoted plant developmental progression with respect to controls in terms of time of flowering and first silique shattering. The days of flowering and first silique shattering following P. liquidambari inoculation were shorted from 34.55 ± 0.33 to 32.50 ± 0.26 and 51.75 ± 0.38 to 47.45 ± 0.35, respectively (Figure 2b and 2c). Meanwhile, P. liquidambari‐inoculated plants showed similar rosette leaves at bolting and total leaves

2.8

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Statistical analyses

at flowering when compared with controls (Figure 2d and 2e). The inflorescence number was increased by P. liquidambari at 40 dai

All experiments were carried out at least three times. The data are pre-

(Figure 2f). Silique production was also measured as an index of fertil-

sented as mean with standard error (SE). Data were subjected to anal-

ity and plant fitness. A positive effect of P. liquidambari was observed

ysis of variance via Tukey's multiple comparison method or Student's t

in siliques number at 45 dai and silique yield that increased by 1.37‐

test using SPSS software (version 18.0) to determine the significant

and 1.11‐fold, respectively (Figure 2g and 2h).

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FIGURE 1 Colonization pattern of Phomopsis liquidambari in Arabidopsis thaliana roots. (a) Root colonization by P. liquidambari at 12 dai. (b‐k) Root colonization by P. liquidambari at 20 dai. (b) Runner hyphae on the root surface. (c, d, and f) Runner hyphae penetrated the epidermal cells through forming thicker bulbous invasive hyphae on the root surface (White *). (g) Branched hyphopodia‐like infection structures during P. liquidambari infection. (h) Phomopsis liquidambari infection through crack sites of the root surface. (i) Penetration of P. liquidambari into the root cortex. (j and k) The lateral root was crowded with abundant hyphae. Bars: (a, e) 50 μm, (b, c, d, f, g, h, and i) 25 μm, (j and k) 100 μm. dai = days after inoculation. [Colour figure can be viewed at wileyonlinelibrary.com]

To determine whether P. liquidambari inoculation shortens

quantification methods. Absolute and relative quantification showed

flowering time through regulating flowering pathway, the expression

similar fungal colonization dynamics (Figure 3b and S2). At 4 dai, we

of key flowering time regulators, including FLC, FT, SOC1, and LFY in

could already detect the presence of the P. liquidambari‐specific ITS

rosette leaves was analysed. During the flowering pathway of

gene in the roots (Figure 3b). After that, the fungal concentration

Arabidopsis, the MADS‐box gene FLC is the major repressor of plant

was increased rapidly with the growth of the plant and reached its

flowering through negatively regulating the expression of floral

peak at 20 dai and then it decreased (Figure 3b). The period over

integrators, such as FT, SOC1, and LFY (Dotto, Gómez, Soto, & Casati,

which the initial fungal concentration declined corresponds to the

2018; Putterill & Varkonyi‐Gasic, 2016). Using RT‐qPCR with

time of flower opening of Arabidopsis. At 20 dai, almost no floral buds

independent samples, we found a slight but not significant difference

were open, whereas 3.75 floral buds on average were open at 24 dai

in FLC expression following P. liquidambari inoculation (Figure 2i). By

(Figure 3b). At the end of the experiment, we rarely detected the

contrast, floral integrators FT, SOC1, and LFY were significantly

presence of the fungal‐specific ITS in plant roots (Figure 3b). We then

stimulated in plants exposed to P. liquidambari (Figure 2i). Taken

collected primary root (0.5 cm) at 20 and 24 dai and processed for

together, these results suggest that P. liquidambari acts as a mutualis-

microscopic observation. More green fluorescent hyphae were found

tic fungus of Arabidopsis and promotes its growth, flowering,

on the root surface of Arabidopsis at 20 dai than that at 24 dai (Figure

and yield.

3c). Meanwhile, we determined the fungal concentration efficiency by using GFP. Similarly, the fungal concentration reached its peak at 20 dai and then it decreased at 24 dai (Figure 3d).

3.3 | Flowering restricts Phomopsis liquidambari colonization in Arabidopsis

To further verify that flowering negatively affects P. liquidambari root colonization, the flowering mutant lines, flc and lfy, were used. The flc and lfy lines showed flowering acceleration and delay,

We sampled Arabidopsis roots and extracted plant genomic DNA to

respectively (Figure 3e and 3f). In spite of flowering time alterations,

assess fungal colonization level with both absolute and relative

the colonization degree of P. liquidambari in root was rapidly increased

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FIGURE 2 Phomopsis liquidambari promotes growth, development, and yield of Arabidopsis thaliana. Effects of P. liquidambari on time of bolting (a), flowering (b), first silique shattering (c) and number of rosette leaves at bolting (d), total leaves at flowering (e), inflorescences at 40 dai (f), siliques at 45 dai (g), and silique yield (h). (i) Effects of P. liquidambari inoculation on expression of flowering time genes. The boxplot represents the data from four independent replicates. The expression of target genes in mock plants was set to 1. The asterisks indicate significant differences between mock and P. liquidambari‐inoculated plants (*p <.05, **p <.01, ***p <.001, t test). Representative seedlings at 12 dai from control (j) and P. liquidambari inoculated treatment (k). E+ = P. liquidambari inoculation; dai = days after inoculation. [Colour figure can be viewed at wileyonlinelibrary.com] after inoculation and decreased at flowering in flc and lfy lines (Figure

flower opening of Arabidopsis. At 16 dai, almost no floral buds were

3e and 3f).

open, whereas 6.45 floral buds on average were open at 20 dai (Figure

Next, we altered flowering time through manipulating ambient temperature. After inoculation, half of Arabidopsis seedlings were

S3). Taken together, these results suggest that it is the flowering that restricts P. liquidambari root colonization.

transferred to another growth chamber with high ambient temperature (27°C). Compared with the plants grown under 23°C, high temperature shortened the flowering time (Figure S3). We found that fungal concentration was increased rapidly with the growth of the

3.4 | Phomopsis liquidambari colonization activates root jasmonate signalling of Arabidopsis at flowering

plant and reached its peak at 16 dai and then it decreased when the ambient temperature was 27°C (Figure S3). The period over which

To test whether jasmonates are involved in flowering‐mediated fungal

the initial fungal concentration declined corresponds to the time of

colonization restriction, we first detected the concentrations of JA and

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FIGURE 3 Colonization dynamics of Phomopsis liquidambari in Arabidopsis thaliana roots. (a) Different developmental stages of A. thaliana flowers. Flowers at stage 13‐14 were counted. (b) The concentration of P. liquidambari in A. thaliana roots at different time points after inoculation. (c) Hyphae assembled on the roots of A. thaliana at 20 and 24 dai. (d) The concentration of P. liquidambari in A. thaliana roots at different time points after inoculation when employing GFP. (e) The concentration of P. liquidambari in flc and lfy A. thaliana roots at different time points after inoculation. The boxplot represents the data from four independent replicates. The colonization level of P. liquidambari at 4 dai was set to 1. dai = days after inoculation. [Colour figure can be viewed at wileyonlinelibrary.com]

its bioactive JA‐Ile at flowering (24 dai). Our results showed that the

(Figure 4c‐4f). Moreover, root mutualisms have been demonstrated

contents of JA and JA‐Ile were higher in P. liquidambari‐inoculated

to induce systemic responses of plant aerial parts (Adolfsson et al.,

roots than those in controls at anthesis (Figure 4a and 4b). Next,

2017; Gerlach et al., 2015), we also detected the JA and JA‐Ile con-

we analysed the expression of JA synthetic and inducible genes,

tents, and the expression of JA synthetic and inducible genes in the

including Allene Oxide Cyclase 3 (AOC3), Lipoxygenase 3 (LOX3), 12‐

rosette leaves of Arabidopsis. By contrast, similar contents of JA and

Oxophytodienoic Acid Reductase (OPR3), and JA‐activity marker MYC2

JA‐Ile, and the expression of JA synthetic and inducible genes, were

(Wasternack & Song, 2016; Zavala‐Gonzalez et al., 2017). In addition

found in rosette leaves of control and P. liquidambari‐inoculated plants

to LOX3, the expression of JA synthetic and responsive genes in roots

(Figure S4). To determine whether the root jasmonate signalling

was stimulated to a higher degree when exposed to P. liquidambari

was specially induced at flowering, we collected root samples of

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FIGURE 4 Jasmonate concentrations and response of Arabidopsis thaliana roots are enhanced by Phomopsis liquidambari at flowering. Analysis of JA (a) and JA‐Ile (b) concentrations in roots of A. thaliana at flowering (24 days after fungal inoculation). Roots of A. thaliana at flowering were collected for JA and JA‐Ile analysis. Bars are means ± SE of five independent replicates. Expression of AOC3 (c), LOX3 (d), OPR3 (e), and MYC2 (f) in roots of mock and P. liquidambari‐inoculated plants at flowering. Bars are means ± SE of four independent replicates. The expression of target genes in mock plants was set to 1. The asterisks indicate significant differences between mock and P. liquidambari‐inoculated plants (*p <.05, **p <.01, t test). E+ = P. liquidambari inoculation. [Colour figure can be viewed at wileyonlinelibrary.com]

Arabidopsis at seedling and rosette stages and detected jasmonate signalling. Our results showed that jasmonate signalling were also activated at seedling stage and then decreased to uninoculated control

3.5 | Jasmonate signalling negatively affects root glucose and fructose concentrations and P. liquidambari colonization

at rosette stage (Figure S5 and S6). In addition to jasmonate, ethylene is another key signal that affects

Given that root jasmonate signalling is induced at flowering upon P.

plant growth and development and mediates plant–microbe interac-

liquidambari inoculation and that jasmonates are important regulators

tions (Camehl et al., 2010; Zhang et al., 2018). Additionally, flowering

of plant–fungal symbioses and carbohydrate partitioning and metabo-

has also been reported to modulated ethylene levels (Diezel et al.,

lism (Gerlach et al., 2015; Hause & Schaarschmidt, 2009; Landgraf

2011). Therefore, we analysed ethylene signalling of Arabidopsis upon

et al., 2012; Machado et al., 2013, 2015; Zavala‐Gonzalez et al.,

P. liquidambari inoculation at flowering. Because ethylene levels in

2017), we hypothesized that flowering may restrict P. liquidambari col-

Arabidopsis were difficult to measure when plants were cultured in

onization through jasmonate‐dependent carbohydrate partitioning

growth substrate, transgenic seedlings expressing ethylene‐responsive

and metabolism alterations. To test this hypothesis, we profiled root

marker EBS:GUS (β‐glucosidase) and ethylene biosynthesis reporter

starch, soluble sugars, soluble protein, and fungal colonization of WT

ACS7:GUS were used. Similar EBS:GUS and ACS7:GUS expressions

and jasmonate perception‐deficient lines (coi1‐2) across different

and activities in leaves and roots were found in P. liquidambari‐inocu-

developmental stages. COI1 (coronatine insensitive 1) encodes an F‐

lated plants and controls at flowering (Figure S7), suggesting that eth-

box protein as a JA‐Ile receptor and mediates core jasmonate signal-

ylene did not response to P. liquidambari at flowering. Together, these

ling and responses (Xie, Feys, James, Nieto‐Rostro, & Turner, 1998).

data demonstrated that root jasmonate contents and signalling were

Except for stage, the treatment and genotype regimes did not affect

induced at the onset of flowering upon P. liquidambari inoculation.

root starch levels in WT and coi1‐2 plants (Figure S8a). In terms of root

ZHANG

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ET AL.

soluble sugars, the levels of sucrose showed significant differences in

5c). Glucose and fructose in WT plants, but not coi1‐2, were

stage and treatment. Compared with WT, coi1‐2 plants showed higher

decreased following P. liquidambari inoculation at the flowering stage

sucrose at early mature stage (Figure 5a). Meanwhile, the presence of

(Figure 5b and 5c). Whether inoculated or not, root soluble protein

P. liquidambari increased sucrose content in both WT and coi1‐2 plants

levels of WT were similar to those observed in coi1‐2 plants across

at the seedling stage (Figure 5a). The variation patterns of root glucose

different developmental stages (Figure S8b). Root starch, sucrose,

and fructose contents were similar with significant differences in

glucose, and fructose contents varied between developmental stages

treatments, genotypes, and developmental stages (Figure 5b and 5c).

(Figure 5a‐5c and S8a). These results suggested that jasmonates

The coi1‐2 plants contained higher glucose and fructose than those

reduce the constitutive and fungus‐induced accumulation of glucose

in WT plants at flowering and early mature stages (Figure 5b and

and fructose at the flowering stage.

FIGURE 5 Effects of Phomopsis liquidambari inoculation on root soluble sugar concentrations of WT and jasmonate‐insensitive (coi1‐2) Arabidopsis thaliana plants across different developmental stages. Average (± SE) sucrose (a), glucose (b), and fructose (c) concentrations in A. thaliana plants. (d) Concentrations of P. liquidambari in WT and coi1‐2 A. thaliana roots across different developmental stages. The colonization level of P. liquidambari in WT plants was set to 1. Bars are means ± SE of five independent replicates. Different letters indicate significant differences within developmental stages (p <.05, one‐way analysis of variance (ANOVA) with Tukey's test). Asterisks indicate significant differences in fungal concentration between genotypes (WT and coi1‐2) within developmental stages (*p <.05, *** p <.001, t test). E+ = P. liquidambari inoculation. [Colour figure can be viewed at wileyonlinelibrary.com]

10

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ET AL.

Moreover, we detected the colonization extent of P. liquidambari in

analysed relationship between root soluble sugar concentrations and

WT and coi1‐2 plants across different developmental stages. In line with

fungal biomass at onset of anthesis. The sugar transport and metabo-

our hypothesis, we found that coi1‐2 plants contained higher fungal fre-

lism mutant plants showed different root sucrose, glucose, and fruc-

quency than that in WT plants at the flowering and early mature stages

tose concentrations at flowering following P. liquidambari inoculation

(Figure 5d). Consistently, the colonization degree of P. liquidambari in

(Figure 6a and 6b). Meanwhile, we found that sucrose concentration

myc2‐2 plants was high. A slight but not significant difference in P.

was not correlated with fungal biomass (p =.0923; Figure 6c). By

liquidambari colonization was found between WT and 35Spro:MYC2

contrast, an obvious positive correlation was observed between the

plants (Figure S9). This may be due to that 35Spro:MYC2 plants do not

glucose and fructose contents and fungal biomass across different

show constitutive activation of JA responses (Chen et al., 2011). It is

sugar transport and metabolism genotypes (p <.001; Figure 6d).

worth noting that fungal colonization degree in coi1‐2 plants was lower

Moreover, an in vitro plate experiment with various combinations of

than that in WT plants at seedling stage (Figure 5d). Additionally, no

sucrose, glucose, and fructose showed that glucose and fructose

significant difference in fungal colonization level was found between

rather than sucrose increased P. liquidambari growth (Figure S10 and

WT and coi1‐2 plants at the rosette stages (Figure 5d).

S11). Collectively, these data suggest a positive effect of root glucose and fructose on fungal frequency within the root.

3.6 | Phomopsis liquidambari colonization levels are positively correlated with root glucose and fructose across different jasmonate genotypes at flowering

3.7 | Jasmonate‐dependent soluble sugar depletion is associated with decreased phloem sugar transport and root invertase activity

To determine whether root glucose and fructose affect P. liquidambari colonization, we used different sugar transport (sweet11, sweet12,

Over the course of the day, jasmonate‐perception deficient coi1‐2

suc1, and suc2) and metabolism (cinv1 and cwinv2) mutant lines and

plants showed more sucrose at 11:00, and (Figure 7a) more glucose

FIGURE 6 Glucose and fructose positively influence P. liquidambari colonization in Arabidopsis thaliana root at flowering. Concentrations of sucrose (a) and glucose + fructose (b) in different A. thaliana sugar metabolism (cinv1 and cwinv2) and sugar transport (sweet11, sweet12, suc1, and suc2) mutant plant roots after P. liquidambari inoculation at flowering. Correlation between sucrose (c) or glucose + fructose (d) and fungal colonization. Detection of soluble sugars and fungal concentration was performed in five independent replicates. Different letters indicate significant differences within different sugar metabolism and transport mutant lines (p <.05, one‐way analysis of variance with Tukey's test). [Colour figure can be viewed at wileyonlinelibrary.com]

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ET AL.

FIGURE 7 Diurnal jasmonate‐dependent reductions of soluble sugars and soluble invertase activity in Phomopsis liquidambari‐inoculated roots. Average (± SE) sucrose (a), glucose + fructose (b), soluble invertase (c), and cell wall invertase (d) in mock and P. liquidambari‐inoculated roots of wild‐type and jasmonates insensitive (coi1‐2) plants across different time points within a day. Bars are means ± SE of five independent replicates. Correlation between sucrose: glucose + fructose and soluble (f) and cell wall (h) invertase activity. Different letters indicate significant differences within each time point (p <.05, one‐way analysis of variance (ANOVA) with Tukey's test). E+ = P. liquidambari inoculation. [Colour figure can be viewed at wileyonlinelibrary.com]

and fructose at 18:00 and 22:00 than WT (Figure 7b). When inoculated

the seedlings were inoculated or not, there were no significant differ-

with P. liquidambari, coi1‐2 plants accumulated sucrose similar to WT,

ences in cell wall invertase activity between WT and coi1‐2 (Figure

and more glucose and fructose from 08:00 to 22:00 (Figure 7a and 7b).

7d). Moreover, we profiled root soluble sugars and invertase and

Next, we detected the activity of root soluble invertase and cell

analysed their correlation relationships. We found a significant

wall invertase in WT and coi1‐2 plants in the presence or absence of

negative correlation between Suc‐to‐Glu+Fru ratios and soluble

P. liquidambari at flowering. Without the P. liquidambari, the activity

invertase activity (p <.001; Figure 7e). No correlation relationship

of soluble invertase in coi1‐2 was higher than that in WT at 08:00

was observed between Suc‐to‐Glu+Fru ratios and cell wall invertase

and 14:00 to 22:00 (Figure 7c). Meanwhile, in the presence of fungus,

activity (p =.571; Figure 7f).

soluble activity of the invertase of coi1‐2 was higher with respect to

Moreover, we sampled phloem sap and detected the concentra-

WT from 11:00 to 22:00 (Figure 7c). By contrast, no matter whether

tions of soluble sugars. As shown in Figure 8a, the concentrations of

12

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ET AL.

FIGURE 8 Diurnal jasmonate‐dependent reductions of phloem sap soluble sugars and sugar transporter activity in Phomopsis liquidambari‐ inoculated plants. Average (± SE) phloem sap sucrose content (a), phloem sap glucose + fructose content (b), leaf sugar transporter activity (c), and root sugar transporter activity (d) in mock and P. liquidambari‐inoculated plants of wild‐type and jasmonate insensitive (coi1‐2) lines across different time points within a day. Bars are means ± SE of five independent replicates. Different letters indicate significant differences among different treatments within each time point or each gene (p <.05, one‐way analysis of variance with Tukey's test). E+ = P. liquidambari inoculation. [Colour figure can be viewed at wileyonlinelibrary.com] sucrose in the phloem sap were higher in the coi1‐2 plants than WT at

4

|

DISCUSSION

08:00 to 18:00. Similarly, upon fungal inoculation, the concentrations of sucrose in the phloem sap are higher in the coi1‐2 plants than WT

Root beneficial fungi have been demonstrated to induce precocious

at 08:00 and 22:00 (Figure 8a). In terms of glucose and fructose, their

flowering of multiple plant species under laboratory and field condi-

concentrations in phloem sap of coi1‐2 were higher than those in WT

tions (Das et al., 2012; Pan et al., 2017; Zavala‐Gonzalez et al.,

from 08:00 to 22:00, and in phloem sap of P. liquidambari‐inoculated

2017); however, little is known about how flowering affects root‐

coi1‐2 plants were higher than those in P. liquidambari‐inoculated

fungal mutualisms. Herein, we report that flowering induces root

WT at 11:00, 18:00, and 22:00 (Figure 8b). Meanwhile, we detected

jasmonate signalling and thus reduces root glucose and fructose

the expression of the main sugar transporters in the rosette leaves

concentrations upon P. liquidambari inoculation and these reductions

and roots at 14:00 (Durand et al., 2018). The expression of SWEET11

negatively affect P. liquidambari root colonization.

and SWEET12 in WT rosette leaves was inhibited following fungal

The generalist root endophyte P. liquidambari establishes mutualis-

inoculation at flowering, and impaired jasmonate signalling alleviated

tic associations with monocotyledonous and dicotyledonous plants,

the negative effect of P. liquidambari on SWEET12 expression (Figure

including A. thaliana (Zhang et al., 2018; Zhou et al., 2018), which is

8c). Meanwhile, fungal inoculation inhibited the expression of SUC1

known as nonhost for ectomycorrhiza and arbuscular mycorrhiza

in the roots of WT rather than coi1‐2 (Figure 8d).

(Almario et al., 2017). The detailed information about the fungal colo-

Taken together, these results suggest that activation of jasmonate

nization strategy is a prerequisite to study plant–fungal interactions

signalling of P. liquidambari‐inoculated plants by flowering negatively

(Su et al., 2013). A large proportion of P. liquidambari hyphae are con-

affects phloem sugar transport and root soluble invertase activity,

fined to root surface, epidermis, and cortex, not extending to the root

which may contribute to the reductions of root glucose and fructose

steles and shoots, suggesting that host plants limit its spread within

concentrations.

roots in a space and quantity pattern (Zhou et al., 2017). Squeezed

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ET AL.

or breached plant cells during P. liquidambari infection indicate the

derived signals that function in the root (Landgraf et al., 2012). It has

presence of turgor pressure and plant cell wall degradation enzymes

been reported that high levels of JA and JA‐Ile were accumulated in

that facilitate fungal spread into the roots (Zuccaro et al., 2011). This

the flowers of N. attenuata as a flower‐specific defence (Li, Wang,

is consistent with recent transcriptome analyses that plant cell wall

et al., 2017). Therefore, root jasmonates of P. liquidambari‐inoculated

hydrolysis‐related enzymes of P. liquidambari, such as the glycoside

plants might be partly originally from the flowers. Further experiments

hydrolase family, were activated during the mutualistic stage (Zhou

with genetic manipulation of flowering time and/or jasmonate signal-

et al., 2017). Moreover, this colonization strategy morphologically

ling pathway together with analysis of phloem sap will help to better

reflected the broad host compatibility of P. liquidambari (Zuccaro,

understand how flowering activates root jasmonate signalling

Lahrmann, & Langen, 2014), as similar infection structures were previ-

following P. liquidambari inoculation. Combined with the results that

ously observed during the infection of rice and peanut by P.

plant deficiency in jasmonate perception (coi1‐2) exhibited higher

liquidambari (Zhang et al., 2018; Zhou et al., 2017; Zhou et al., 2018).

colonization levels of P. liquidambari at flowering, we proposed that

However, compared with rice‐P. liquidambari interaction, we found

jasmonates, acting as mediators, are involved in flowering‐mediated

notable differences in colonization patterns. Although the colonization

P. liquidambari root colonization limitation. It is noteworthy that the

is restricted, a fraction of P. liquidambari hyphae enter the central

root jasmonate signalling was also induced at seedling stage after P.

cylinder of rice roots and spreading systemically to shoots through

liquidambari inoculation. A similar situation has also been reported in

vascular tissue (Zhou et al., 2018). The possible explanation for the

Arabidopsis–P. indica interaction and mycorrhizal association (Hause

host‐adapted colonization strategy of P. liquidambari is the coevolution

& Schaarschmidt, 2009; Lahrmann et al., 2015; Pozo, López‐Ráez,

of plant and fungal partners (Zhou et al., 2018; Zuccaro et al., 2014).

Azcón‐Aguilar, & García‐Garrido, 2015). This early jasmonate response

Plant developmental stages have been demonstrated to influence

might function as positive regulator during the colonization of P.

the rhizosphere fungal and bacterial community assembly and root

liquidambari, as coi1‐2 plants exhibited low fungal colonization than

exudates act as chemical basis driving this microbiota change

that in WT. Jasmonates have been reported to enhance root‐fungal

(Chaparro, Badri, & Vivanco, 2014; Zhalnina et al., 2018), but the

symbiosis through induction of flavonoids, reorganization of cytoskel-

knowledge of plant development influences on root microbial commu-

eton, and enhancement of plant fitness (Genre & Bonfante, 1998;

nity, especially fungal mutualism is missing (Lundberg et al., 2012).

Hause & Schaarschmidt, 2009; Steinkellner et al., 2007). Flavonoids

Across different developmental stages, we used both absolute and rel-

are JA‐inducible and stimulate germination and growth of fungi and

ative quantification to evaluate the fungal colonization efficiency and

act as signals between symbiotic partners (Steinkellner et al., 2007).

found that it is the flowering that restricted colonization of P.

Reorganization of microtubular cytoskeletion is required for the estab-

liquidambari in Arabidopsis roots. When we altered flowers opening

lishment of fungal symbiosis, which is in response to jasmonates

time through using flowering mutants and manipulating ambient tem-

(Koda, 1997; Genre & Bonfante, 1998). Thus, jasmonates play dual

perature, the fungal colonization was still decreased at flowering,

roles in root‐P. liquidambari mutualisms, depending on symbiotic stage

which further confirmed this notion. Similar results were obtained by

(Pozo et al., 2015). At the early stage, jasmonates orchestrate plant

Luo et al. (2017), who reported that the colonization rate of arbuscular

response to coordinate host plants for recognition and establishment

mycorrhizal fungi in the rice root was increased from tillering to

of symbiosis. When the symbiosis is fully established, the function of

jointing stage and then decreased at the flowering and ripening stages.

jasmonates switches to restrict fungal spread. Interestingly, WT and

These results raised a possibility that flowering might stimulate a

coi1‐2 plants showed similar fungal colonization level at rosette stage,

systemic response that drives underground fungal cooperation loss.

suggesting that the early jasmonate response was not required for

Root jasmonate signalling of P. liquidambari‐inoculated plants was

entire life of P. liquidambari within root. This study combined with

activated at the onset of flowering, suggesting the possibility that

our previous study, which showed that auxin signalling is required

flowering and limitation of fungal colonization are linked by

for the establishment of root‐P. liquidambari symbiosis (Zhang et al.,

jasmonates. It is unknown how this kind of root jasmonate signalling

2018), suggesting the possible involvement of a phytohormone

was established. It is tempting to speculate that flowering producing

crosstalk during P. liquidambari symbiosis.

secondary signals, such as other phytohormones, peptides, or electri-

The results described above raised the question of how jasmonate

cal activity, upon P. liquidambari inoculation, spread to the P.

signalling is linked with flowering‐mediated P. liquidambari root coloni-

liquidambari‐inoculated roots, where they activate jasmonate signal-

zation decline. Because jasmonates have been proposed to be

ling. For instance, indoleacetic acid acts as a secondary signal in

involved in plant–fungal interactions through alterations of sink carbo-

response to herbivore attacks, and it rapidly moves to systemic tissues

hydrate status (Gerlach et al., 2015; Landgraf et al., 2012; Zavala‐

and thereby activates jasmonates and jasmonate‐dependent plant

Gonzalez et al., 2017), we analysed root starch, soluble sugars, and

defence (Machado et al., 2016). Moreover, peptides have been

protein of WT and jasmonate‐insensitive plants across different devel-

reported to be important mobile messengers that complement the

opmental stages upon P. liquidambari inoculation. Root glucose and

regulatory function of phytohormones (Takahashi & Shinozaki,

fructose contents were dramatically reduced in P. liquidambari‐inocu-

2019). Meanwhile, jasmonates themselves could act as mobile signals

lated plants at flowering, suggesting that hosts provided less carbohy-

that spread to distant parts (Schulze et al., 2019). During the autoreg-

drate to the fungal partner. Floral organs are able to perform

ulation of nodulation, jasmonates have been proposed as shoot‐

photosynthesis because green petals have chlorophylls and Rubisco

14

ZHANG

enzymes, but they become fully heterotrophic once opening (Borghi &

ET AL.

suggested a starvation‐mediated fungal colonization limitation, sugar

Fernie, 2017; Thomas et al., 2003). Carbohydrates and amino acids

signalling‐mediated plant resistance might be involved in the regula-

support the flowers' growth and metabolism, such as formation of col-

tion of fungal colonization (Bezrutczyk et al., 2018). Sugar concentra-

our and scent (Muhlemann et al., 2014; Borghi & Fernie, 2017). Thus,

tions have been demonstrated to be modulated during pathogen

flowering is an energy consuming process and reevaluates the symbi-

infection and trigger signalling cascades, which induce plant defence

otic quality based on cost–benefit trade‐offs and sanctions against

gene (Gebauer et al., 2017). Furthermore, previous studies have deter-

costly symbionts. Furthermore, plants tend to require more nitrogen

mined that the restriction of fungal symbiont was due to the accumu-

at reproductive stage, and most of mutualistic fungi facilitate the host

lation of secondary metabolites, especially glucosinolates, which are

to uptake phosphate (Almario et al., 2017; Ezawa & Saito, 2018),

induced by jasmonates (Hiruma et al., 2016; Lahrmann et al., 2015).

which may decrease the dependency of plants on fungal symbionts.

Thus, we could not exclude the possibility that plant secondary metab-

Flowering‐mediated root P. liquidambari colonization restriction was

olites play a role in flowering‐mediated restriction of P. liquidambari

probably because of cost is over benefit. Reduced root symbiont fre-

colonization. Additionally, fungal growth is not just dependent on

quency could save carbohydrate to ensure the success of flowering

sugars. It is conceivable that some other essential nutrients, such as

and subsequent seed set. Over colonization of fungal symbionts,

fatty acids, amino acids, and organic acids, are associated with

which is caused by intense inoculation or compromised plant

flowering‐mediated fungal colonization reductions (Bezrutczyk et al.,

innate immunity, would negatively affect plant growth, development,

2018; Jiang et al., 2017), as such metabolites are also modulated at

and reproduction, such as flowering and sterility (Zavala‐Gonzalez

flowering (Borghi & Fernie, 2017).

et al., 2017).

The carbohydrate strength of sink tissue is primarily determined by

Based on the following observations, we speculate that jasmonate‐

the ability of phloem loading/unloading that is strongly affected by the

dependent root glucose and fructose depletion drive the decline of P.

activity of sugar transporters and metabolism (Durand et al., 2018;

liquidambari root colonization at flowering. First, jasmonate signalling

Kühn & Grof, 2010). Thus, we are interested in whether the

deficiency prevented the reductions in root glucose and fructose con-

jasmonate‐mediated root soluble sugars decrease at flowering is

centrations, and fungal colonization of P. liquidambari‐inoculated

through alterations of source‐sink sugar transport and root invertase.

plants at flowering. Second, glucose and fructose rather than sucrose

Over the course of the day, whether inoculation or not, a deficiency

are the preferred C resources for fungal partners. Compared with con-

in jasmonate perception increased root soluble sugar contents, sugar

trol and sucrose, glucose and fructose increased P. liquidambari growth

transport, and root soluble invertase activity of Arabidopsis plants dur-

using MSM medium supplemented with various combinations of solu-

ing the light phase, suggesting that jasmonate‐dependent reductions

ble sugars. Consistent to this result, Sun et al. (2019) demonstrated

in constitutive and fungus‐induced accumulation of root glucose and

that glucose and fructose are the preferred C resources for P.

fructose were likely the consequence of both repressions of sugar

liquidambari based on in vitro and in vivo experiments. In a given eco-

transport and root invertase. Similar results were obtained by

system, microbial substrate preferences and nutrient availability deter-

Machado et al. (2015), who reported that N. attenuata irAOC plants,

mine the abundance of specific microbes (Zhalnina et al., 2018).

which are impaired in jasmonate biosynthesis display higher levels of

Microbial metabolite substrate preference could be reflected in their

glucose and fructose, as well as soluble invertase activity. Unlike

genomic and transcriptomic traits. The existence of acid intracellular

pathogen infection, which is often associated with cell wall‐bound

invertase tvinv in the genome of Trichoderma virens enables the fungus

invertase (Essmann et al., 2008; Proels & Hückelhoven, 2015), root

to hydrolyse and use sucrose. When tvinv is knocked out, the fungal

soluble invertase are produced in response to P. liquidambari, reflecting

germling preferably feeds on glucose over sucrose (Vargas et al.,

the different strategies in nutrient acquisition between beneficial and

2009). Piriformospora indica acquires glucose from symbiotic interface

pathogenic fungi. Invertases have been demonstrated to determine

through hexose transporter (Rani, Raj, Dayaman, Kumar, & Johri,

the ratio of glucose and fructose to sucrose (Bhaskar et al., 2010;

2016). Substrate preferences of P. liquidambari on glucose and fruc-

Jin, Ni, & Ruan, 2009; Machado et al., 2015). Bhaskar et al. (2010)

tose over sucrose suggesting three processes that existed may involve

reported that silencing a potato (Solanum tuberosum) vacuolar acid

in the access of fungus to photosynthates. (a) Sugars are transported

invertases (Vinv) gene increased sucrose but reduced glucose and

from shoots to roots by SWEETS/SUC proteins. (b) Sucrose is hydro-

fructose concentrations. It is noteworthy that glucose and fructose

lyzed to glucose and fructose by root invertases. (c) Glucose and fruc-

contents are not always negatively correlated with sucrose concentra-

tose are transported into fungal hyphae. Further works with dissection

tion, as other factors, such as photosynthetic activity and phloem

of the genome and transcriptome would help to target the partners

sugar transport, regulate the ratios of Suc‐to‐Glu+Fru (Machado

that are involved in nutrients exchange at the symbiotic interface of

et al., 2015). CO2 is fixed in mesophyll cells and then transported into

P. liquidambari and Arabidopsis. Moreover, analysis of root soluble

other C‐demand organs by phloem loading/unloading with facilitation

sugars and fungal colonization with different sugar transport and

of sugar transporters (Durand et al., 2018). Analysis of phloem sap in

metabolism mutant lines revealed a positive correlation between P.

WT and jasmonate perception impaired plants with or without P.

liquidambari colonization and glucose+fructose rather than sucrose,

liquidambari inoculation showed that phloem sugar transport plays a

further confirm the links of glucose and fructose concentrations and

role in jasmonate‐mediated declines of root soluble sugars at

P. liquidambari colonization degree. Although the above results

flowering. Previous studies with poplar and N. attenuata have

ZHANG

15

ET AL.

demonstrated that jasmonates have the ability to regulate carbohy-

experiments. CCD supervised all work. All of authors read and

drate transport (Babst et al., 2005; Machado et al., 2015). The expres-

approved the final manuscript.

sion of the main sugar transporters further confirmed this notion. Invertases and phloem sugar transport are often coupled to regulate

ORCID

the sink sugar pool (Doidy et al., 2012; Naseem, Kunz, & Dandekar,

Chuan‐Chao Dai

https://orcid.org/0000-0002-0980-0018

2017). Sink invertases have been demonstrated to positively influence the activity of phloem sugar transport (Doidy et al., 2012). Because

RE FE RE NC ES

jasmonate signalling was induced in P. liquidambari‐inoculated‐roots

Adolfsson, L., Nziengui, H., Abreu, I. N., Šimura, J., Beebo, A., Herdean, A., … Spetea, C. (2017). Enhanced secondary‐ and hormone metabolism in leaves of arbuscular mycorrhizal Medicago truncatula. Plant Physiology, 175, 392–411. https://doi.org/10.1104/pp.16.01509

rather than leaves, a negative feedback on phloem sugar transport might be produced by the inhibited root soluble invertase. Further studies with root‐specific inactivation of jasmonate signalling and invertases are required to reveal the mechanisms behind the cooperation between root soluble invertases and sugar transport in root glucose and fructose reductions upon P. liquidambari inoculation at flowering. Taken together, our results revealed a novel metabolic network in which plant flowering activates root jasmonate signalling, which then reduces concentrations of root glucose and fructose upon beneficial fungus P. liquidambari inoculation. The declines in root soluble sugars are probably due to the inhibitions of phloem sugar transport and root soluble invertase activity and restrict root colonization of P. liquidambari. Given that the present study was conducted in greenhouse with single plant and fungal species, and the outcomes of plant‐beneficial fungal interactions are dynamic, varying on plant and fungal species, resource availability, and environmental conditions (Johnson et al., 1997; Nelson, Hauser, Hinson, & Shaw, 2018; Saikkonen, Wäli, Helander, & Faeth, 2004), further studies will be required to investigate the prevalence of flowering‐mediated root fungal cooperation loss in the context of various fungal colonization modes,

plant

resources,

and

environmental

conditions.

Our

results highlight importance of plant flowering in root microbiota composition. ACKNOWLEDGEMEN TS The authors are grateful for Zi‐Qiang Zhu (Nanjing Normal University), Wei‐Ming Shi, Guang‐Jie Li (Institute of Soil Science, China Academy of Science), and Ning‐Ning Wang (Nankai University) for sharing mutants and transgenic lines of Arabidopsis seeds. The authors thank Kai‐Min Hui of Instrumental Analysis Centre for her help with the microscopic imaging. The research was financially supported by National Natural Science Foundation of China (NSFC 31870478), the Priority Academic Program Development of Jiangsu Higher Education Institutions, and the Doctor Breeding Project of Nanjing Normal University (1812000006317). The authors express their great thanks to anonymous reviewers and editorial staff for their time and attention. CONF LICT OF INT E RE ST The authors declare no conflict of interest. AUTHOR CONTRIBUTIONS WZ designed the experiments, analyse the data, and wrote and revised the manuscript. WZ, JY, TC, MJT, KS, SLS, and FJX performed

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Zavala‐Gonzalez, E. A., Rodríguez‐Cazorla, E., Escudero, N., Aranda‐Martinez, A., Martínez‐Laborda, A., Ramírez‐Lepe, M., … Lopez‐Llorca, L. V. (2017). Arabidopsis thaliana root colonization by the nematophagous fungus Pochonia chlamydosporia is modulated by jasmonate signaling and leads to accelerated flowering and improved yield. New Phytologist, 213, 351–364. https://doi.org/10.1111/nph.14106 Zhalnina, K., Louie, K. B., Hao, Z., Mansoori, N., da Rocha, U. N., Shi, S., … Brodie, E. L. (2018). Dynamic root exudate chemistry and microbial substrate preferences drive patterns in rhizosphere microbial community assembly. Nature Microbiology, 3, 470–480. https://doi.org/ 10.1038/s41564‐018‐0129‐3 Zhang, W., Sun, K., Shi, R. H., Yuan, J., Wang, X. J., & Dai, C. C. (2018). Auxin signalling of Arachis hypogaea activated by colonization of mutualistic fungus Phomopsis liquidambari enhances nodulation and N2‐ fixation. Plant, Cell & Environment, 41, 2093–2108. Zhang, W., Wang, H. W., Wang, X. X., Xie, X. G., Siddikee, M. A., Xu, R. S., & Dai, C. C. (2016). Enhanced nodulation of peanut when co‐inoculated with fungal endophyte Phomopsis liquidambari and bradyrhizobium. Plant Physiology and Biochemistry, 98, 1–11. https://doi.org/10.1016/ j.plaphy.2015.11.002 Zhang, W., Wang, X. X., Yang, Z., Ashaduzzaman, S. M., Kong, M. J., Lu, L. Y., … Dai, C. C. (2017). Physiological mechanisms behind endophytic fungus Phomopsis liquidambari‐mediated symbiosis enhancement of peanut in a monocropping system. Plant and Soil, 416, 1–18. https:// doi.org/10.1007/s11104‐017‐3183‐3 Zhou, J., Li, X., Chen, Y., & Dai, C. C. (2017). De novo Transcriptome assembly of Phomopsis liquidambari provides insights into genes associated with different lifestyles in rice (Oryza sativa L.). Frontiers in Plant Science, 8, 121. Zhou, J., Li, X., Huang, P. W., & Dai, C. C. (2018). Endophytism or saprophytism: Decoding the lifestyle transition of the generalist fungus Phomopsis liquidambari. Microbiological Research, 206, 99–112. https:// doi.org/10.1016/j.micres.2017.10.005 Zhu, Z., An, F., Feng, Y., Li, P., Xue, L., Mu, A., … Guo, H. (2011). Derepression of ethylene‐stabilized transcription factors (EIN3/EIL1) mediates jasmonate and ethylene signaling synergy in Arabidopsis. Proceedings of the National Academy of Sciences, USA, 108, 12539–12544. Zuccaro, A., Lahrmann, U., Güldener, U., Langen, G., Pfiffi, S., Biedenkopf, D., … Kogel, K. H. (2011). Endophytic life strategies decoded by genome and transcriptome analyses of the mutualistic root symbiont Piriformospora indica. PLoS Pathogens, 7, e1002290. https://doi.org/ 10.1371/journal.ppat.1002290 Zuccaro, A., Lahrmann, U., & Langen, G. (2014). Broad compatibility in fungal root symbioses. Current Opinion in Plant Biology, 20, 135–145. https://doi.org/10.1016/j.pbi.2014.05.013

SUPPORTING INFORMATION Additional supporting information may be found online in the Supporting Information section at the end of the article. Figure S1. Phomopsis liquidambari inoculation increases chlorophyll content. Twelve days after P. liquidambari inoculation, Arabidopsis thaliana rosette leaves were collected for chlorophyll content analysis. Bars are means ± SE of four independent experiments. Asterisks indicate significant differences in chlorophyll content between treatments (mock and P. liquidambari inoculation; *P < 0.05, t test). E+ = P. liquidambari inoculation. Figure S2. Colonization dynamics of Phomopsis liquidambari in Arabidopsis thaliana roots. (a) The standard curve was obtained by

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the Ct values and the logarithm of the known concentration of five‐

Arabidopsis rosette leaves and roots were sampled at flowering (24

point ten folds dilution of DNA. (b) The concentration of P.

days after fungal inoculation) for EBS:GUS (a) and ACS7:GUS (b) assays.

liquidambari in A. thaliana roots at different time points after inocula-

(c) The relative GUS activity of EBS:GUS and ACS7:GUS in rosette

tion. The colonization level of P. liquidambari was measured by abso-

leaves and roots of Arabidopsis at flowering. E+ = P. liquidambari

lute quantification method with P. liquidambari specific ITS gene.

inoculation.

Figure S3. Effects of different ambient temperature (23 oC and 27 oC)

Figure S8. Effects of Phomopsis liquidambari inoculation on root starch

on root colonization dynamics of P. liquidambari. The boxplot repre-

and soluble protein of wild‐type (WT) and jasmonate‐insensitive (coi1‐

sents the data from four independent replicates. The colonization level

2) Arabidopsis thaliana plants across different developmental stages.

of P. liquidambari at 4 dai was set to 1. dai, days after inoculation.

Average (± SE) starch (a) and soluble protein (b) concentrations in

Figure S4. Phomopsis liquidambari inoculation does not affect jasmonates concentrations or response of Arabidopsis thaliana rosette leaves at flowering. Analysis of JA (a) and JA‐Ile (b) concentrations in leaves of A. thaliana at flowering (24 days after fungal inoculation).

WT and coi1‐2 Arabidopsis thaliana roots. Bars are means ± SE of five independent replicates. Different letters indicate significant differences within developmental stages (P < 0.05, one‐way ANOVA with Tukey's test). E+ = P. liquidambari inoculation.

Bars are means ± SE of five independent replicates. Expression of

Figure S9. Colonization degree of P. liquidambari in different

AOC3 (c), LOX3 (d), OPR3 (e) and MYC2 (f) in rosette leaves of mock

Arabidopsis thaliana jasmonate signaling lines at flowering. Bars are

and P. liquidambari‐inoculated plants at flowering. Bars are means ±

means ± SE of four independent replicates. Different letters indicate

SE of four independent replicates. The expression of target genes in

significant differences within different jasmonate signaling lines (P <

mock plants was set to 1. E+ = P. liquidambari inoculation.

0.05, one‐way ANOVA with Tukey's test).

Figure S5. Phomopsis liquidambari inoculation enhances jasmonates

Figure S10. Glucose and fructose increase Phomopsis liquidambari

concentrations and response of Arabidopsis thaliana roots at seedling

growth. Average diameter (± SE) of P. liquidambari on MSM medium

stage. Analysis of JA (a) and JA‐Ile (b) concentrations in roots of A.

with different combinations of soluble sugars at 4 and 7 days. Bars

thaliana at seedling stage (8 days after fungal inoculation). Bars are

are means ± SE of three independent experiments. Different letters

means ± SE of five independent replicates. Expression of AOC3 (c),

indicate significant differences among treatments at 7 days (P <

LOX3 (d), OPR3 (e) and MYC2 (f) in roots of mock and P.

0.05, one‐way ANOVA with Tukey's test).

liquidambari‐inoculated plants at seedling stage. Bars are means ± SE

Figure S11. Effects of different concentrations (mg L‐1) of sucrose, glu-

of four independent replicates. The expression of target genes in mock plants was set to 1. Asterisks indicate significant differences between mock and P. liquidambari‐inoculated plants (*P < 0.05, t test). E+ = P. liquidambari inoculation. Figure S6. Phomopsis liquidambari inoculation does not affect jasmonates concentrations or response of Arabidopsis thaliana roots at rosette stage. Analysis of JA (a) and JA‐Ile (b) concentrations in

cose and fructose applications on the phenotype of Phomopsis liquidambari. P. liquidambari was grown on MSM medium with different combinations of soluble sugars for 7 days and collected for phenotype analysis under a DIC microscope. Bars: 10 μm. Table. S1 List of primers used in the study (Khatabi et al., 2012; Zavala‐Gonzalez et al., 2017)

roots of A. thaliana at rosette stage (16 days after fungal inoculation). Bars are means ± SE of five independent replicates. Expression of

How to cite this article: Zhang W, Yuan J, Cheng T, et al.

AOC3 (c), LOX3 (d), OPR3 (e) and MYC2 (f) in roots of mock and P.

Flowering‐mediated root‐fungus symbiosis loss is related to

liquidambari‐inoculated plants at rosette stage. Bars are means ± SE

jasmonate‐dependent root soluble sugar deprivation. Plant Cell

of four independent replicates. The expression of target genes in mock

Environ. 2019;1–19. https://doi.org/10.1111/pce.13636

plants was set to 1. E+ = P. liquidambari inoculation. Figure S7. Phomopsis liquidambari inoculation does not affect ethylene signaling of Arabidopsis rosette leaves and roots at flowering.