Engineered coumarin accumulation reduces mycotoxin‐induced oxidative stress and disease susceptibility

Summary

Coumarins can fight pathogens and are thus promising for crop protection. Their biosynthesis, however, has not yet been engineered in crops. We tailored the constitutive accumulation of coumarins in transgenic Nicotiana benthamiana, Glycine max and Arabidopsis thaliana plants, as well as in Nicotiana tabacum BY‐2 suspension cells. We did so by overexpressing A. thaliana feruloyl‐CoA 6‐hydroxylase 1 (AtF6’H1), encoding the key enzyme of scopoletin biosynthesis. Besides scopoletin and its glucoside scopolin, esculin at low level was the only other coumarin detected in transgenic cells. Mechanical damage of scopolin‐accumulating tissue led to a swift release of scopoletin, presumably from the scopolin pool. High scopolin levels in A. thaliana roots coincided with reduced susceptibility to the root‐parasitic nematode Heterodera schachtii. In addition, transgenic soybean plants were more tolerant to the soil‐borne pathogenic fungus Fusarium virguliforme. Because mycotoxin‐induced accumulation of reactive oxygen species and cell death were reduced in the AtF6’H1‐overexpressors, the weaker sensitivity to F. virguliforme may be caused by attenuated oxidative damage of coumarin‐hyperaccumulating cells. Together, engineered coumarin accumulation is promising for enhanced disease resilience of crops.

Introduction

Soybean (Glycine max) is a rich source of oil, amino acids, and protein and thus belongs to the economically most important crops on Earth (FAOSTAT, 2019; Hartman et al., 2011). The USA, Brazil and Argentina are the major soybean‐producing countries providing more than 80% of the global soybean yield (FAOSTAT, 2019). Several diseases threaten soybean production and frequently result in economic losses (Bandara et al., 2020; Wrather et al., 2010). For example, in Argentina and the USA, the sudden death syndrome (SDS), which is caused by the soil‐borne fungus Fusarium virguliforme, is a frequent late‐season disease of the soybean crop. The fungus infects the root of susceptible seedlings and then invades their vascular tissue. It does, however, not seem to infest aerial parts of the plant (Roy et al., 1997; Rupe, 1989). Foliar symptoms appear with a huge delay, often as late as at the beginning of flowering or even later (Roy et al., 1997). Disease symptoms comprise interveinal chlorosis and necrosis and, eventually, premature defoliation. The latter is caused by mycotoxins that translocate from the infected root to the aerial parts of the plant (Brar et al., 2011; Chang et al., 2016). In Argentina, 14% of the soybean yield loss have been assigned to SDS (Wrather et al., 2010) and in the USA, between 1996 and 2016, an estimated economic loss of almost 4000 USD per hectare was ascribed to SDS (Bandara et al., 2020). These losses classify SDS as one of the most threatening soybean diseases. Measures to control SDS are limited because SDS‐resistant soybean cultivars are unavailable and fungicides often ineffective (Westphal, 2008). SDS frequently coincides with the occurrence of soybean cyst nematodes (SCN; Heterodera glycines), raising a huge economic bill to soybean production in the USA and the South Americas (Bandara et al., 2020; Koenning and Wrather, 2010; Wrather et al., 2010). When H. glycines and F. virguliforme occur together, SDS often is particularly detrimental (McLean and Lawrence, 1993; Westphal et al., 2014; Xing and Westphal, 2006). Thus, generating soybean varieties with dual resilience to F. virguliforme and H. glycines represents a promising strategy to securing soybean yield.

Scopoletin is a compound in the coumarin class of secondary metabolites and a potent antioxidant (Beyer et al., 2019). Because of its antifungal, antibacterial, antiviral, and insect deterrent activity, the coumarin qualifies for providing broad‐spectrum pest and disease resistance (Beyer et al., 2019; Churngchow and Rattarasarn, 2001; Fay and Duke, 1977; Garcia et al., 1995; Gómez‐Vásquez et al., 2004; Goy et al., 1993; Olson et al., 1991; Prats et al., 2002, 2006; Stringlis et al., 2018; Sun et al., 2014; Tal and Robeson, 1986; Tripathi et al., 2011; Valle et al., 1997). Consistently, the endogenous level of scopoletin frequently correlates with the extent of disease susceptibility or resistance in various tobacco cultivars and hybrids, sunflowers and various species in the Platanus and Hevea genera (Chong et al., 2002; El Modafar et al., 1995; El Oirdi et al., 2010; Garcia et al., 1999; Goy et al., 1993; Prats et al., 2007; Sun et al., 2014). The extent of scopoletin accumulation depends on the genotype, age, organ and physiological condition of the plant. For example, in the roots of Arabidopsis thaliana (subsequently referred to as Arabidopsis in this paper), scopoletin presence is constitutive. Further biosynthesis and secretion to the rhizosphere are induced during iron deficiency (Fourcroy et al., 2014; Schmid et al., 2014; Siwinska et al., 2014, 2018) or upon root colonization by beneficial rhizobacteria that trigger an induced systemic resistance (ISR) response (Stringlis et al., 2018). Secretion of simple coumarins contribute to plant tolerance to iron starvation (Fourcroy et al., 2014; Schmid et al., 2014; Schmidt et al., 2014), induces shifts in the composition of synthetic root microbiome communities (Voges et al., 2019) and selectively suppresses soil‐borne pathogens while promoting the multiplication of ISR‐inducing microbes (Stringlis et al., 2018). By doing so, scopoletin and other coumarins are thought to modulate the composition of the root microbiome to the benefit of the plant. Aerial parts of healthy Arabidopsis plants do not constitutively accumulate scopoletin. In leaves, de novo biosynthesis of scopoletin is rather induced by abiotic (e.g. low temperature, 2,4‐dichlorophenoxyacetic acid) and biotic stress (e.g. F. oxysporum, Phakopsora pachyrhizi) (Beyer et al., 2019; Döll et al., 2018; Kai et al., 2006). Different from Arabidopsis, soybean leaves do not contain scopoletin even upon inoculation with P. pachyrhizi (Beyer et al., 2019).

In Arabidopsis, scopoletin biosynthesis is associated with expression of the FERULOYL‐CoA 6‐HYDROXYLASE 1 (F6’H1) gene (Beyer et al., 2019; Kai et al., 2008; Schmid et al., 2014). AtF6’H1 encodes a 2‐oxoglutarate‐dependent dioxygenase catalysing the ortho‐hydroxylation of feruloyl‐CoA to 6‐hydroxyferuloyl‐CoA (Kai et al., 2008). Despite inactivity of recombinant F6’H1 towards the esculetin precursor caffeoyl‐CoA and several other potential phenylpropanoid substrates, esculetin and esculin accumulation in the roots of Arabidopsis wild‐type plants has been reported to depend on F6’H1 (Ziegler et al., 2017). In light‐protected organs, such as roots, the isomerization and lactonization of feruloyl‐CoA depends on coumarin synthase (COSY) (Vanholme et al., 2019). In planta, scopoletin may be further metabolized, e.g. to fraxetin and sideretin (Rajniak et al., 2018; Siwinska et al., 2018) or it is glycosylated by UDP‐glycosyltransferases (UGT's) (Chong et al., 2002; Hino et al., 1982; Lim et al., 2003). Scopolin is the glucoconjugate of scopoletin and predominantly accumulates in the vacuole (Taguchi et al., 2000). Different from scopoletin, scopolin has no noticeable antioxidant activity and does not inhibit the intracellular accumulation of reactive oxygen species (ROS; Beyer et al., 2019). It also does not inhibit the germination of, for example, P. pachyrhizi spores (Beyer et al., 2019). However, scopolin inhibits the formation of germ tubes of Sclerotinia sclerotiorum and halts hyphal growth of Alternaria alternata at doses similar to those of active scopoletin (Li and Wu, 2016; Prats et al., 2006). These findings indicate that scopolin may also exert a direct protective function, at least in some plant–fungus interactions. Alternatively, or in addition, scopolin pools in the vacuole may serve as a reservoir for scopoletin that may quickly be released by scopolin‐hydrolysing β‐glucosidases after cellular collapse, e.g. after attack from microbial pathogens, nematodes, or chewing insects. Apart from its direct plant protective properties, scopoletin has been suggested to indirectly contribute to plant disease resistance by suppressing the biosynthesis of fungal virulence factors such as mycotoxins (Gnonlonfin et al., 2011). Increased scopoletin content may also augment the nutraceutical value of plant‐derived products due to its manifold beneficial properties to human health which include its antidiabetic, anti‐inflammatory, anticancer, and anti‐obesity activity (Chang et al., 2015; Ding et al., 2008; Ham et al., 2016; Li et al., 2015).

In this work, we explored the feasibility of AtF6’H1‐overexpression for constitutive accumulation of simple coumarins, particularly scopoletin and scopolin, in transiently transformed Nicotiana benthamiana and in stably transformed Arabidopsis and soybean plants, and in tobacco BY‐2 culture cells. We also evaluated the possible effect of AtF6’H1‐overexpression on various plant–pathogen interactions. We show that homologous and heterologous overexpression of AtF6’H1 mainly leads to constitutive, but differing, accumulation of scopoletin and scopolin between species, organs and compartments. An elevated level of coumarin glycosides in Arabidopsis roots enhanced the resistance to H. schachtii. Constitutive accumulation of coumarins in soybean increased the tolerance to SDS associated with increased capacity of AtF6’H1‐overexpressing cells for buffering excessive mycotoxin‐induced ROS accumulation. Engineered coumarin biosynthesis may, therefore, be utilized for increasing the antioxidant capacity and thereby reducing the stress susceptibility of crops.

Results

Heterologous and transient overexpression of AtF6’H1 in N. benthamiana causes scopoletin accumulation

As the first step towards understanding whether expression of AtF6’H1 suffices for gaining coumarin biosynthesis in another plant species, we transiently overexpressed AtF6’H1 and different N‐ or C‐terminal fusions of the protein with GFP (green fluorescent protein), RFP (red fluorescent protein) or FLAG in N. benthamiana. We then analysed scopoletin abundance, scopoletin fluorescence in UV light, transgene expression and fusion protein accumulation and localization in transformed leaves. Independent of the N‐ or C‐terminus of F6’H1 being translationally fused to a protein tag, transient overexpression of AtF6’H1 caused strong accumulation of scopoletin in N. benthamiana (Figure 1a). Quantitative RT‐PCR, immunoblotting analysis and/or fluorescence detection confirmed the expression of AtF6’H1 and its fusion DNA constructs only in leaves transformed with the respective transgene‐encoding construct but not in p19‐expressing controls (Figure 1b,d–f). In UV light, AtF6’H1‐overexpressing and scopoletin‐accumulating leaves displayed fluorescence, whereas control construct‐transformed leaves expressing GFP, RFP or the silencing inhibitor p19 alone were neither fluorescent nor did they synthesize detectable amounts of scopoletin (Figure 1a,c). Depending on the DNA construct used, scopoletin levels in AtF6’H1‐expressing leaves varied from 0.23 to 1 μg/g FW at 3 days after transformation (Figure 1a). Confocal microscopy of leaves transiently co‐overexpressing AtF6’H1‐GFP and RFP confirmed the predicted localization of F6’H1 (Softberry.com; ProtComp Version 9.0: cytoplasmic with a score of 8.2) to the cytoplasm (Figure 1d).

Figure 1.

Transient overexpression of AtF6‘H1 causes scopoletin accumulation in N. benthamiana. Scopoletin (a) and AtF6‘H1 mRNA transcript (b) in N. benthamiana leaves overexpressing AtF6‘H1 or C‐ or N‐terminal fusions of AtF6‘H1 with GFP, RFP or FLAG controlled by the constitutive CaMV 35S promoter. P19 silencing inhibitor was co‐expressed in all cases. Expression of P19 only served as negative control. Shown are means ± SD of three independent experiments at 3 days after transient transformation. (c) Photos of transiently transformed leaves with characteristic fluorescence of scopoletin (in UV light; upper panel), GFP (middle panel) and RFP (lower panel). (d) AtF6‘H1‐GFP co‐localizes with cytoplasmic RFP in transiently transformed epidermal cells of N. benthamiana. GFP and RFP fluorescence were analysed by confocal laser scanning microscopy. (e, f) Accumulation of GFP and RFP and fusions of both fluorescent proteins and FLAG with AtF6‘H1 was verified by SDS‐PAGE and immunodetection using antibodies specific to GFP, RFP and FLAG.

Stable overexpression of AtF6’H1 causes constitutive but varying spatial accumulation of scopoletin and scopolin among different organs of Arabidopsis and soybean

Next, we analysed the accumulation of coumarins in independent transgenic events of Arabidopsis (accession Col‐0) and soybean (cultivar W82) stably overexpressing AtF6’H1 or FLAG‐AtF6’H1, respectively. Among simple coumarins other than scopoletin and scopolin, we only detected low concentrations of esculin in roots and leaves of transgenic plants (10–100‐fold less than scopolin, Figure S2). We therefore focused our analysis on scopoletin and scopolin. Accumulation of the coumarins and fluorescence of leaves were similar in the different transgenic events of a same species. However, pronounced differences were seen between transgenic Arabidopsis and soybean. Arabidopsis leaves displayed strong scopoletin fluorescence in UV light no matter whether the adaxial or abaxial side of the leaf was inspected (Figure 2a). In contrast, only the abaxial side of transgenic soybean leaves displayed strong fluorescence in UV light, whereas the fluorescence of the upper leaf side was only slightly enhanced when compared to the untransformed controls (Figure 2b). In both Arabidopsis and soybean, fluorescence was most intense in leaf veins and trichomes (Figures 2b and S1). Consistently, the scopolin concentration was up to 5‐fold greater in leaves of Arabidopsis than soybean with a maximum of ~370 μg/g FW and ~80 μg/g FW, respectively (Figure 2c). The scopoletin content was similar among all analysed leaves of transgenic Arabidopsis and soybean events with average levels of ~3 μg/g FW. The identity of both scopoletin and scopolin was confirmed by HPLC (high performance liquid chromatography) and GC–MS (gas chromatography–mass spectrometry) analysis (Figure S2). No or only little scopoletin and scopolin were detected in leaves of untransformed Arabidopsis and soybean. In roots of AtF6’H1‐overexpressing Arabidopsis, scopoletin (8–9 μg/g FW) and scopolin (440–490 μg/g FW) levels were much higher than in roots of transgenic soybean (~0.5 μg/g FW scopoletin and ~25 μg/g FW scopolin) (Figure 2c). Consistent with the generally higher coumarin abundance in transgenic Arabidopsis than soybean leaves and roots, scopoletin and scopolin levels of Arabidopsis seeds also exceeded those in soybean seeds by ~10–20‐fold (Figure S3). To our surprise, the distribution of scopoletin and scopolin in aboveground and belowground organs of transgenic plants of both species was opposite. Scopoletin and scopolin levels were much higher in roots than in the shoot of Arabidopsis, whereas in soybean, the level of the two coumarins was greater in the shoot than in the root (Figure 2c). Neither in Arabidopsis nor soybean any obvious alterations of phenotype due to AtF6’H1 overexpression was seen regarding plant growth and development (Figure S4).

Figure 2.

Stable overexpression of AtF6‘H1 in Arabidopsis and soybean results in constitutive coumarin accumulation. Representative photos of adaxial and abaxial sides of detached leaves of Arabidopsis (a) and soybean (b) wild‐type and AtF6‘H1‐overexpressing transgenic lines were inspected in ambient or UV light. Only the transgenic plants showed characteristic blue fluorescence of leaves in UV light. (c) Scopoletin and scopolin content in roots and leaves of wild‐type Arabidopsis (left panel) and soybean (right panel) and two independent events overexpressing AtF6’H1, respectively. Coumarin concentration in methanolic extracts of 3‐to‐6‐week‐old plants was quantified by HPLC. Shown is the average + SD of three independent experiments.

Total scopoletin content is highest within cells

We next used N. tabacum BY‐2 suspension cells stably overexpressing AtF6’H1 to analyse the cellular distribution of scopoletin and scopolin. In contrast to untransformed BY‐2 culture cells, transgenic cells, and their growth medium both displayed characteristic blue fluorescence in UV light (Figure 3a). As in organs of intact plants, in transgenic BY‐2 cells, scopoletin predominantly accumulated in its glycosylated scopolin derivative. Approximately 77% of the total scopoletin (free and glycosylated) of culture cells were attributed to scopolin which exclusively accumulated inside the cells (Figure 3b). We never detected scopolin in the culture medium of cells. In contrast, scopoletin was more abundant in the medium than in the cells (Figure 3b).

Figure 3.

Scopoletin and scopolin mainly accumulate within the cells. (a) AtF6‘H1‐expressing tobacco BY‐2 cells, but not the untransformed wild type, and their culture medium fluoresce in UV light. Tobacco BY‐2 cells were separated from the growth medium by vacuum filtration at 7 days after subculture. Cells and medium individually inspected in ambient and UV light. (b) Scopoletin and scopolin content of AtF6‘H1‐overexpressing BY‐2 cells and their culture medium relative to the total amount of scopoletin and scopolin in the batch culture. Average values and SD of three technical replicates are shown.

Upon cellular damage, scopolin is quickly hydrolysed to scopoletin

To quantify the time required for the conversion of supposedly vacuole‐localized scopolin to scopoletin by glucosidases (Ahn et al., 2010), we analysed the kinetics of scopolin hydrolysis upon decompartmentalization by simply grinding the leaves. Tissue homogenization is making scopolin available as a substrate to, for example, beta‐glucosidase. While scopolin accounted for almost 90% of the total scopoletin content in undamaged soybean leaves, its content quickly decreased to nearly 50% within 1 h after grinding. The scopoletin content increased correspondingly (Figure 4a). In Arabidopsis, however, the initial concentration of scopolin made up 97% of the total scopoletin content. Of the scopolin content in Arabidopsis leaves, 29% was converted to scopoletin within 1 h after tissue grinding the tissue (Figure 4b). Thereafter, scopolin hydrolysis to scopoletin slowed down. At 24‐h post‐tissue disruption (hpt), more than ~60% and 90% of the initial scopolin had been hydrolysed to scopoletin in the samples from Arabidopsis and soybean, respectively (Figure 4a,b). Thus, upon cellular damage intracellular scopolin seems to be quickly converted to scopoletin.

Figure 4.

Upon destruction of AtF6‘H1‐overexpressing leaves scopolin is hydrolysed to scopoletin. Leaves of transgenic soybean (a) and Arabidopsis (b) plants were homogenized and analysed for their content of scopolin and scopoletin at the indicated times after tissue disruption (hpt). Average values + SD of three biological replicates are shown.

Scopolin hyperaccumulation in Arabidopsis reduces susceptibility to H. schachtii

Because scopoletin and scopolin support plant health and as their concentration was highest in the roots of transgenic Arabidopsis plants, we analysed whether transgenic plants over‐accumulating the two coumarins would better ward of root pathogens. Thus, we investigated the extent of susceptibility of the Arabidopsis wild type, f6’h1 mutant and F6’H1 overexpressor to the root‐attacking beet cyst nematode (BCN) H. schachtii. We found that the number of developed female nematodes at the roots seemed to be similar among all genotypes tested when these accumulated <200 μg scopolin per gram root FW. Infection rates on roots containing <200 μg/g scopolin ranged from 2.5 to 17 females per plant (Figure 5a). Within this group of plants, the f6’h1 mutant which did not accumulate detectable amounts of scopolin in the root seemed to be more susceptible than the wild type with varying amounts of scopolin ranging from 2 to 15 μg/g FW. Strikingly, an apparent and statistically significant reduction in the infection rate of female and male nematodes was seen for the F6’H1 overexpressor with more than 200 μg scopolin per gram root FW (Figures 5b and S5). Thus, increased levels of coumarin glycosides in Arabidopsis roots coincided with reduced susceptibility to H. schachtii. Because low amounts of esculin were detectable in roots of transgenic Arabidopsis plants (Figure S2), other coumarins may also contribute to the observed phenotype.

Figure 5.

Transgenic Arabidopsis plants with high scopolin levels in root are less susceptible to H. schachtii. Wild‐type Arabidopsis, f6‘h1 mutants, AtF6‘H1 overexpressors and azygous Arabidopsis plants were grown on agar plates. At 12 days after inoculation with nematodes infection, severity and root content of scopolin were determined. High scopolin levels in roots exceeding 200 μg/g FW coincided with reduced female numbers. Shown are (a) the root scopolin content and infection scores from individual plates with two plants per genotype, respectively. The average infection severity (b) of groups containing more (group >200) or less (group <200) than 200 μg/g FW scopolin in the root. The asterisk marks statistically significant differences between groups >200 and <200 (t‐test; P < 0.05). Three independent experiments with at least four plates (2 plants per plate) per genotype were performed. Data from (a and b) are derived from identical experiments but illustrated in different way.

Overexpression of AtF6’H1 induces the SDS tolerance of soybean

Next, we evaluated the interaction of untransformed and AtF6’H1‐overexpressing soybean plants with F. virguliforme. Before inoculation, no obvious difference was seen between the roots of the two genotypes of plant. Fourteen days after inoculation, the root of both non‐transgenic and transgenic plants showed rotting indicating successful F. virguliforme infection (Figure 6a). The infected roots were brownish, less dense, and they apparently produced less biomass than non‐infected roots (Figure S6). However, no obvious phenotypic differences were observed between roots of non‐transgenic and AtF6’H1‐overexpressing plants (Figure S6). F. virguliforme infection of roots severely affected shoot biomass of both genotypes as well (Figure 6b). However, the reduction of shoot biomass was significantly lower for the two independent AtF6’H1‐transgenic soybean events (~40% decrease) when compared to the wild type (~50% decrease). To analyse, whether the reduced SDS severity of the transgenic plants resulted from diminished root colonization, we determined the abundance of F. virguliforme DNA in infected roots. Although, fungal DNA abundance was slightly lower in F. virguliforme‐infected roots of transgenic plants compared to wild type, the differences were not statistically significant (Figure 6c). Consistent with these findings, scopoletin in vitro did not significantly inhibit F. virguliforme mycelial growth at concentrations lower than 500 μM (Figure S7). These results suggest that overexpressing AtF6’H1 does not provide direct protection of soybean from F. virguliforme but rather attenuates the damage of SDS on the plant. Constitutive coumarin accumulation in soybean thus seems to induce soybean tolerance to SDS disease.

Figure 6.

Stable overexpression of AtF6‘H1 in soybean induces SDS tolerance. Wild‐type plants and two independent AtF6‘H1‐overexpressing soybean events were hydroponically grown for 7 days prior to root inoculation with spores of F. virguliforme. (a) Wild‐type and transgenic plants show similar root rotting at 14 days after inoculation. (b) Shoot biomass (fresh weight) of infected wild‐type plants or transgenic plants was calculated in comparison with mock‐inoculated plants 14 days after inoculation. Shown are min‐to‐max boxplots with all data points, average values (+) and SD of three independent experiments with nine plants per genotype and treatment. Asterisks indicate significant differences to the wild‐type control according to Dunnett's multiple comparisons test with P ≤ 0.05. (c) Abundance of F. virguliforme DNA in infected roots of transgenic and wild‐type plants at 14 days after inoculation. DNA was extracted from ≥22 plants per genotype for fungal DNA quantification by qRT‐PCR (real‐time quantitative reverse transcription PCR). Shown are min‐to‐max boxplots with all data points, average values (+) and SD of three independent experiments. Differences among wild‐type and transgenic plants were not significant according to Dunnett's multiple comparisons test.

Coumarin accumulation in AtF6’H1‐overexpressing cells attenuates mycotoxin‐induced ROS accumulation and cell death

During infection, many Fusarium species produce mycotoxins that may cause oxidative stress and eventually provoke host cell death (Albrecht et al., 1998; Jennings et al., 2001; Singh and Upadhyay, 2014; Yekkour et al., 2015). One such mycotoxin is fusaric acid (FA; 5‐butylpicolinic acid). In tomato, the toxin promotes ROS production, lipid peroxidation and cell death (Singh and Upadhyay, 2014). In vitro, scopoletin scavenges ROS (Beyer et al., 2019). To see whether coumarins buffer mycotoxin‐induced ROS hyperaccumulation also in planta, we analysed FA‐triggered ROS production in soybean. As shown in Figure 7a, control treatment with water did not notably induce the accumulation of ROS in leaves of wild‐type and AtF6’H1‐transgenic soybean plants. However, a ROS burst was detected in leaves of the wild type after treatment with FA with maximum ROS accumulation at ~25 min post‐treatment (Figure 7b). The ROS response to FA was significantly attenuated in leaves overexpressing AtF6’H1. Maximum ROS levels in transgenic leaves were about half the levels in leaves of the wild type (Figure 7b,c). To validate the effect of AtF6’H1 overexpression on the redox state of plants, we also examined ROS accumulation in non‐transgenic and AtF6’H1‐overexpressing BY‐2 culture cells in response to FA and the Fusarium toxin deoxynivalenol (DON). DON was previously reported to trigger ROS production and cell death in BY‐2 cells (Yekkour et al., 2015). Figure 7d reveals that both FA and DON induced a distinct ROS signal in wild‐type BY‐2 cells. The accumulation of ROS was more enhanced upon treatment with DON (6‐fold) than with FA (2‐fold) compared to water‐treated cells. Neither FA nor DON elicited ROS accumulation in AtF6’H1‐transgenic, scopoletin‐accumulating BY‐2 cells (Figure 7d). Consistently, mycotoxin‐induced cell death was reduced in transgenic BY‐2 cells when compared to the non‐transgenic cells (Figure 7e). While the ratio of living to dead wild‐type cells decreased enormously after treatment with FA or DON, the viability of transgenic cells was not significantly affected by either of the two mycotoxins (Figure 7f). Together, these data suggest that increasing the content of coumarins in transgenic cells increases the tolerance to mycotoxin‐induced oxidative stress and by doing so seems to prevent ROS‐induced cell damage and death.

Figure 7.

AtF6‘H1 overexpression is associated with decreased ROS accumulation upon mycotoxin exposure. Leaf discs from trifoliate leaves of 19‐day‐old wild‐type or transgenic soybean were floated on (a) water or (b) 30 μg/mL FA. ROS accumulation was recorded over time by luminol‐amplified chemiluminescence. Average values ± SD from three independent experiments are shown. (c) Area under the curve (AUC) values (mean ± SEM) were calculated from relative luminescence intensities (RLUs) as measured by the ROS assay for (a) water and (b) FA treatment. Asterisks mark statistical difference relative to the respective treatment in wild‐type cells (Sidak's multiple comparisons test, P < 0.0001). (d) Treatment with FA and DON induces ROS production in wild‐type BY‐2 cells but not in RFP‐AtF6’H1‐overexpressing cells. Cells were treated with 30 μg/mL FA or 50 μg/mL DON or water 7 days after subculture and ROS levels quantified by measuring fluorescence intensity in a plate reader following addition of the dye 2′,7′‐dichlorofluorescin diacetate (H2DCFDA). Average values and SD from three independent experiments are shown. Asterisks mark statistical difference relative to the respective treatment in wild‐type cells (Sidak's multiple comparisons test, P < 0.0001). (e) Viability of wild‐type and transgenic BY‐2 cells was analysed by confocal laser scanning microscopy 4 h after treatment with FA, DON or water (control) and addition of FDA and propidium iodide. (f) Differences were semi‐quantitively measured by counting pixels in the respective FDA or PI channels and calculating ratios. Shown are average values and SD from three independent experiments. Asterisks mark statistical difference relative to the respective control treatment (Dunnett's multiple comparisons test, P < 0.001).

Discussion

Scopoletin occurs in many plant species (Gnonlonfin et al., 2012) in which it is often involved in the response to biotic and abiotic stress (Beyer et al., 2019; Döll et al., 2018; Kai et al., 2006). However, scopoletin is absent from aerial parts of soybean even in stressful condition (Beyer et al., 2019). Numerous studies reported the successful engineering of metabolic pathways to accumulate phenylpropanoids (e.g. hydroxycinnamates, flavonoids, isoflavonoids, anthocyanins and stilbenes) in various crops including maize (Liu et al., 2018a,b), rice (Ogo et al., 2013; Shin et al., 2006; Zhu et al., 2017), tomato (Zhang et al., 2015), alfalfa (Deavours and Dixon, 2005), tobacco (Hain et al., 1993; Xie et al., 2006) and soybean (Yu et al., 2003). However, to our knowledge there is no study on the successful engineering of coumarin biosynthesis in crops so far. We took advantage of F6’H1 as the exclusive limiting enzyme for mainly producing scopoletin from its substrate feruloyl‐CoA that is widely abundant in plants. Despite recombinant AtF6’H1 showing no significant activity towards any tested substrate other than feruloyl‐CoA in vitro (Kai et al., 2008), esculetin and esculin concentrations are even significantly lower in roots of the f6h1 mutant than in roots of the wild type (Ziegler et al., 2017), suggesting at least some minor affinity of F6’H1 for caffeoyl‐CoA in vivo to produce esculetin and, following subsequent glycosylation, esculin. Stable overexpression of AtF6’H1 resulted in coumarin accumulation in all analysed tissues of the plant species tested (Figures 1 and 2). The lower concentrations of scopoletin and scopolin in transgenic soybean when compared to Arabidopsis plants may be linked to the differential availability of scopoletin's major precursor feruloyl‐CoA in the two species. Similarly, the content of the feruloyl‐CoA precursor ferulic acid considerably varies among plant species (Zhao and Moghadasian, 2008). Likewise, opposite root‐to‐leaf ratios of scopoletin and scopolin in transgenic Arabidopsis (~5:1) and soybean (~1:5) may result from different substrate availability in the organs of the two species, different expression levels of the transgene, varying catalytic activity of F6’H1, or differential movement characteristics of scopoletin in Arabidopsis and soybean. In fact, both acropetal and basipetal long‐distance transport of flavonoids have been reported for Arabidopsis (Buer et al., 2007). Similarly, UV‐induced elicitation of resveratrol biosynthesis caused its accumulation in distal leaves above and below the UV‐challenged leaf, indicating bidirectional transport of the polyphenol in grapevine (Wang et al., 2013). For scopolin, a root‐to‐shoot transport has recently been confirmed in Arabidopsis (Robe et al., 2021). Although unclear whether scopoletin or scopolin is translocated, both have been detected at high concentration in tobacco leaf midveins and Arabidopsis xylem sap (Dieterman et al., 1964; Robe et al., 2021), suggesting that scopoletin may also be transported throughout the plant.

In the analysed transgenic plants and cells, scopoletin levels never exceeded 3–5 μg/g FW but rather accumulated as scopolin (Figures 2 and 3). The distribution of scopoletin and scopolin within transgenic BY‐2 cells and in the culture medium was as described before (Okazaki et al., 1982); scopoletin almost exclusively accumulated outside the cells, whereas scopolin was absent from the outside of the cells but accumulated to high levels within cells (Figure 3). Similarly, high scopolin‐to‐scopoletin ratios (13:1) were recorded in tobacco leaves (Sargent and Skooge, 1960). Due to the potential concentration‐dependent toxicity of scopoletin not only for microbes but also for plant cells (Graña et al., 2017), glycosylation of scopoletin and translocation of scopolin to the vacuole likely serve as a mechanism of scopoletin tolerance. Free extracellular scopoletin is a known substrate of peroxidases that oxidize scopoletin in the presence of H₂O₂ (Chong et al., 1999). This confirms scopoletin as a strong antioxidant not only in vitro (Beyer et al., 2019) but also in vivo. Dysregulation of the redox balance by excessive extracellular scopoletin may harm plant cells and cause fitness penalties to the plant. Thus, the rapid uptake of scopoletin from the medium and subsequent glycosylation likely provides a cellular detoxification mechanism when critical concentrations have been reached (Taguchi et al., 2000). However, plants can benefit from a fast provision of the bioactive scopoletin through rapid hydrolysis of the o‐glycosidic bond of scopolin. In the incompatible interaction of tobacco with tobacco mosaic virus, in which the hypersensitive response leads to necrotic lesions, scopoletin is quickly released and likely reduces ROS formation (Chong et al., 2002). Also, the accumulation of scopoletin coincides with plant infection from necrotizing fungal pathogens, such as Botrytis cinerea (El Oirdi et al., 2010). Hence, a higher pool of scopolin could potentially enhance the resistance to necrotizing pathogens by providing a pool for the fast scopolin hydrolysis to provide scopoletin. This hydrolysis is being catalysed by ER‐localized β‐glucosidases (e.g. Arabidopsis BGLUs 21–23) (Ahn et al., 2010) upon the release of scopolin from the vacuole, e.g. when cells are damaged (Figure 4).

BGLU23, which is also known as PYK10 (Nakano et al., 2017), might be particularly interesting. Its expression is induced upon infection with the BCN H. schachtii (Nitz et al., 2001) of Arabidopsis roots where it could be involved in releasing scopoletin from scopolin to counter‐infection. High levels of scopolin coincided with reduced numbers of both female and male nematodes developing at the root (Figures 5 and S5). Whether this reduction is caused by scopolin or locally by released scopoletin remains to be elucidated. Previous studies suggested a correlation between nematode‐induced ROS accumulation and an increase of antioxidants, such as flavonoids and camalexin (Amjad Ali et al., 2014; Hamamouch et al., 2020; Labudda et al., 2018). Nematodes themselves can also actively modulate plant secondary metabolism, e.g. by secreting chorismate mutase (Vanholme et al., 2009). Yet, to our knowledge, there is no report on the correlation between nematode infection and the accumulation of coumarins in plants. However, recent studies revealed scopoletin to be a potent biocontrol agent against SCN, with scopoletin being active after the treatment of soybean seeds with the compound (Yan et al., 2021). SCNs are in the same genus as H. schachtii and form a disease complex with F. virguliforme, thereby leading to stronger SDS severity of soybean (Westphal et al., 2014; Xing and Westphal, 2006). Increased resistance of coumarin‐accumulating soybean lines to SCN may provide a promising strategy for controlling both SCN and F. virguliforme.

Scopoletin inhibited the growth of F. virguliforme in vitro only at high concentrations (Figure S7). This might, at least in part, explain why no effect on root infection severity was observed for transgenic soybean roots accumulating relatively low levels of scopoletin (Figure 6a,c). Consistent with our results, an IC50 of ~1 mM scopoletin was observed in growth inhibition assays with F. solani (Sacc.) Mart. (Peterson et al., 2003), further highlighting the need for high in planta concentrations to directly target the fungal pathogen. Still, plants hyperaccumulating scopoletin were less affected by F. virguliforme and exhibited less fresh weight reduction than wild‐type plants (Figure 6b).

Foliar symptoms of Fusarium diseases include water loss of leaves coupled to decreased photosynthesis efficiency (Yan et al., 2018). However, fresh weight reduction of leaves upon detaching, indicative for transpiration‐associated water loss, was identical in transgenic lines and wild‐type plants (Figure S8), suggesting that increased coumarin concentrations apparently did not directly affect transpiration rate. Instead AtF6’H1 overexpression may provide tolerance to SDS by other means. Apart from its direct antimicrobial activity, scopoletin has been suggested to also indirectly contribute to plant disease resistance by suppressing production of fungal virulence factors like mycotoxins (Gnonlonfin et al., 2011). Consistent with this finding, increasing the content of antioxidant secondary metabolites in plants has been proposed as a potential strategy to improve plant resistance or tolerance, especially for mycotoxin‐producing fungi such as Fusarium species (Atanasova‐Penichon et al., 2016). The potential of this strategy is further supported by several studies describing the induction of secondary metabolism genes such as those with a role in flavonoid or glyceollin synthesis in response to F. virguliforme infection (Lozovaya et al., 2004; Marquez et al., 2019; Radwan et al., 2011). These often antioxidative metabolites may also be beneficial by improving the capacity of a plant to buffer excessive mycotoxin‐induced ROS production and alleviate the associated cellular damage (Yekkour et al., 2015). Indeed, many studies suggest that F. virguliforme mycotoxins induce ROS that are supposed to damage the tissue through different mechanisms (e.g. Rubisco degradation, membrane degradation and programmed cell death), thereby contributing to foliar symptom development (Abeysekara et al., 2016; Brar et al., 2011; Jin et al., 1996). Increased concentrations of antioxidant coumarins such as scopoletin in plant tissues may therefore contribute to securing plant fitness in stressful conditions by scavenging excessive stress‐induced ROS, thereby minimizing chlorophyll degradation, oxidation of lipids, proteins and carbohydrates or other cellular damage associated with oxidative stress (Yu, 1994). Here, we demonstrated that in both AtF6’H1‐overexpressing soybean plants and BY‐2 cells, mycotoxin‐induced ROS accumulation was reduced compared to the wild type (Figure 7a–d). The reduction of ROS correlated with decreased cell death in transgenic cells (Figure 7e,f) and might explain the increased tolerance of AtF6’H1‐transgenic plants to SDS (Figure 6b). Similarly, scavenged ROS accumulation by coumarins in germ tubes of P. pachyrhizi antagonized the development of fungal pre‐infection structures (Beyer et al., 2019). However, in field trials in Brazil, the susceptibility of AtF6’H1‐overexpressing lines was slightly, but not significantly, reduced compared to the azygous controls (Figure S9). This finding indicates that the accumulation of coumarins in leaves is not sufficient to provide high levels of protection from SBR (Asian soybean rust). Delivery of spore germination‐inhibiting coumarins such as scopoletin to the soybean leaf surface might be needed for an effective coumarin‐mediated defence response to the rust fungus. A blueprint of such a defence strategy exists in sunflower. In this plant, increased coumarin secretion to the phylloplane of certain cultivars correlated with their capacity to suppress germination of spores of the sunflower rust fungus Puccinia helianthi (Prats et al., 2007).

Due to the reported function of scopoletin as an insect feeding deterrent when mixed with artificial diets (Tripathi et al., 2011), we further compared the interactions between larvae of the generalist insect pest S. exigua and soybean wild‐type and F6’H1 overexpressing plants. In a two‐choice feeding bioassay, leaf discs of F6’H1 overexpressing soybean trifoliate leaves were significantly less damaged by feeding larvae than leaves of wild‐type controls (Figure S10), confirming coumarins as an insect repellent in planta and further highlighting their broad‐spectrum plant‐protective potential.

Apart from reducing disease susceptibility, scopoletin may also support the recruitment of beneficial microbes to the rhizosphere, as previously described for Arabidopsis (Lundberg and Teixeira, 2018; Stringlis et al., 2018). Indeed, several reports suggest that an enhanced transcription of F6’H1 orthologs in soybean roots is associated with the colonization by endophytic microbes such as Piriformospora indica and Bradyrhizobium japonicum (Bajaj et al., 2018; Libault et al., 2010). In sum, due to its beneficial properties for plants and humans (e.g. through antidiabetic, anti‐inflammatory, anticancer and anti‐obesity activity) (Chang et al., 2015; Ding et al., 2008; Ham et al., 2016; Li et al., 2015), increasing the concentration of scopoletin and other coumarins in crops may provide a broadly useful agronomic trait. We here showed that engineering this trait is feasible by targeted modulation of the expression of AtF6’H1 in plants.

Experimental procedures

Plants and fungi

Nicotiana benthamiana seeds were grown in ED73‐type soil (Einheitserde Werksverband) in the greenhouse for 4–6 weeks before performing Agrobacterium‐mediated transient transformation.

Arabidopsis seeds were sown on pipette tips filled with 0.5× Hoagland (1 mM KH2PO4, 1 KNO3, 1 mM MgSO4, 5 mM CaNO3; 50 μM H3BO3, 0.05 μM CoCl2, 0.05 μM CuSO4, 15 μM ZnSO4, 2.5 μM KI, 50 μM MnSO4, 3 μM Na2MnO4, 50 μM Fe3+‐EDTA, pH 5) supplemented with agar as previously described (Fourcroy et al., 2014). In short, pipette tips were filled with 0.5× Hoagland supplemented with 0.8% (w/v) agar and placed in a box containing 0.5× Hoagland medium. One seed was placed on each agar‐filled pipette tip allowing roots to grow through the tip into the medium below.

For easier separation of root and upper plant parts, soybean plants were grown in 50‐mL tubes containing 0.5× Hoagland medium. Growth conditions were maintained at 24 °C and a light period of 16 h.

The suspension cell culture of N. tabacum cv. BY‐2 (cultivar Bright Yellow‐2, BY‐2) was grown in a Murashige and Skooge‐based medium (4.3 g/L MS‐medium, 30 g/L Sucrose, 200 mg/L KH₂PO₄, 0.2 mg/L 2.4‐Dichlorophenoxyacetic acid, 1 mg/L Thiamine‐HCL, 100 mg/L Myo‐inositol; adjusted to pH 5.8 with 1 M KOH). Cells were subcultured once a week, by pipetting ~5% of cells into fresh medium.

Fusarium virguliforme (Wa1‐ss1 (Malvick and Bussey, 2008)) was kindly provided by Dean Malvick (University of Minnesota) and kept on potato dextrose agar (Carl Roth) at ambient temperature and in light deprivation. Spores were harvested by lightly scraping the surface soaked with ddH₂O and subsequent filtration through gauze.

Transient transformation of N. benthamiana leaves

A. tumefaciens (AGL01) was used for transient transformation of N. benthamiana leaves. Agrobacteria were grown and resuspended in infiltration buffer (10 mM MgCl₂; 10 mM MES, pH 5.6; 150 μM acetosyringone) to a final OD600 of 1. Agrobacteria containing the silencing suppressor P19 (Voinnet et al., 2003) were grown to a final OD of 0.5 and mixed 1:1 with Agrobacteria containing the gene of interest shortly before being infiltrated into N. benthamiana leaves using a syringe. Plants were further kept under long‐day conditions (16 h light, 8 h dark, 22 °C) for 3 days, to ensure sufficient accumulation of protein. Leaves were then harvested for further analysis.

Pictures of leaves exposed to UV light (Blak‐Ray®, Analytik Jena) were taken using a Sony α 6000 in constant settings. GFP fluorescence was visualized in the ChemiDoc™ MP(BIO‐RAD) with blue epi illumination and the 530/28 emission filter. RFP fluorescence was visualized with green epi illumination and 605/50 emission filter. Equal exposure time for all samples ensured comparability.

Cellular localization of fusion proteins

For the analysis of the localization of GFP and RFP fusion proteins, confocal laser scanning microscopy was performed using the TCS SP8 (Leica). Leaf discs were punched out from transformed N. benthamiana leaves after 3 days. The leaf discs were then placed on a glass slide for further microscopy, where GFP and RFP fluorescence were inspected to analyse the localization of the fusion proteins. For GFP detection, the excitation wavelength of the argon laser was 488 nm and emission monitored at 500–530 nm. RFP was excited with a diode‐pumped solid‐state (DPSS) laser at 561 nm and emission detected at 590–625 nm.

Stable transformation of plants and plant cell culture

DNA vectors for stable transformation of plants were generated using Gateway® Cloning as described by the manufacturer (Thermo Fisher). Primers used in this work are listed in Table S2.

For stable Arabidopsis transformation, the floral dip method was used (Clough and Bent, 1998). Therefore, the coding sequence of AtF6’H1 (AT3G13610) was cloned into the pAMPAT‐GWY (kindly provided by Ralph Panstruga) overexpression vector.

Stably transformed soybean plants were provided by the Transformation Facility of the Iowa State University. The construct used for transformation comprised an omega translational enhancer element, a FLAG‐protein tag to facilitate detection of the protein and the coding sequence of AtF6’H1. The vector pB2GW7 (Karimi et al., 2002) was used as backbone.

BY‐2 cell suspension culture was transformed using an established protocol (Graumann and Evans, 2011). For the stable transformation into wild‐type BY‐2 tobacco cell suspension cells (kindly provided by Fraunhofer IME), the AtF6’H1 was cloned into the DNA vector pSITE‐4NB (Chakrabarty et al., 2007) providing an N‐terminal mRFP‐tag. Three‐day‐old cells were co‐incubated with transformed agrobacteria for 3 days in the dark at 26 °C on modified MS agar plates (see plants and fungi). After incubation, the cells were washed with medium containing 50 mg/mL timentin and subsequently plated on modified MS agar plates containing 50 mg/mL kanamycin for the selection of transformed cells. Cells were then incubated at 26 °C for 4–6 weeks until callus formation.

Real‐time quantitative reverse transcription PCR (qRT‐PCR) analysis

The abundance of AtF6’H1 transcript in transiently transformed N. benthamiana leaves was determined by qRT‐PCR analysis as described (Beyer et al., 2019). Total RNA was extracted from ground leaf material as described (Chomczynski and Sacchi, 1987). RNA was then transcribed to cDNA according to manufacturer's instructions using 9‐mer random oligonucleotides (ThermoFisher Scientific) and RevertAid Reverse Transcriptase (ThermoFisher Scientific). To quantify the transcript abundance, qRT‐PCR was performed as per manufacturer's instructions in a BIO‐RAD CFX384 REAL‐TIME System using iTaq™ Universal SYBR8® Green Supermix (BIO‐RAD) and expression normalized to NbACT2. The primers used for qRT‐PCR are listed in Table S2.

Protein isolation and analysis

Protein was extracted from plant material as previously described (Conrath et al., 1997). Total protein was then fractioned via SDS‐PAGE and subsequently transferred to nitrocellulose membrane for immunodetection (Conrath et al., 1997). Primary monoclonal antibodies against GFP and RFP were obtained from Roche and Chromotek, respectively. Antigen–antibody complexes were detected with horseradish peroxidase‐linked anti‐mouse secondary antibody (Cell Signalling Technology) followed by chemiluminescence detection with Clarity™ Western ECL Substrate (BIO‐RAD) in the ChemiDoc™ (BIO‐RAD). To check for equal loading Ponceau S (AppliChem), staining was conducted following immunodetection.

Scopoletin and scopolin extraction

Plant material was extracted as described (Beyer et al., 2019). In short, plant material was frozen in liquid nitrogen, ground, weighed and extracted in 90% methanol for 16 h. After centrifugation at full speed for 15 min, the supernatant was evaporated in a concentrator plus (Eppendorf) and the pellet resuspended in 300 μL of methanol. Samples were stored at −20 °C until further analysis.

HPLC analysis

For HPLC analysis, 10 μL sample were separated on a c‐18 reverse phase column heated to 40 °C (NUCLEODUR 100‐5 C18 ec, MACHEREY‐NAGEL). Solvents used for separations were water and acetonitrile both supplemented with 1.5% acetic acid. The flow rate was set at 0.8 mL/min with a gradient (Figure S11) beginning at 10% acetonitrile. The fluorescent compounds scopoletin and scopolin, and esculetin and esculin were detected in the fluorescence detector (RF‐20A, Shimadzu) at 365 nm excitation and 470 nm emission and identified by the retention time compared with authentic standards. Retention times were as follows: esculin: ~6.2 min; scopolin: ~6.6 min; esculetin: ~7.9 min; and scopoletin ~11.6 min. Further verification of compound identity was done by inspecting absorption spectra (230–500 nm) in the photodiode array (SPD‐M40, Shimadzu). Scopoletin (Tokyo Chemical Industry Co., Ltd.) and scopolin (Aktin Chemicals, Inc.) content was calculated referring to calibration curves prepared with authentic standards. For the complete set‐up of HPLC components (all from Shimadzu), see Table S1.

Nematode experiments

f6’h1 T‐DNA insertional mutant SALK_132418 (Kai et al., 2008) was obtained from NASC. Homozygous insertion of the T‐DNA in the F6’H1‐gene was confirmed by PCR according to the recommended procedure (http://signal.salk.edu/tdnaprimers.2.html). Seeds of the Arabidopsis wild‐type Col‐0, f6’h1 mutant and F6’H1‐overexpressor plants were surface‐sterilized for 30 s in 70% of ethanol followed by 3 min in 1.2% of sodium hypochlorite. Sterile seeds were washed three times, dried on sterile filter paper and stored at 10 °C to promote stratification. Sowing was performed on Knop agar (Sijmons et al., 1991) and subsequently, Arabidopsis was kept at RT and 16 h photoperiod. After 5 days, the scopolin synthesis of the overexpression line was validated via fluorescence and verified. Non‐expressive, azygous plants were transferred separately onto fresh Knop agar with two plants per plate. In order to keep conditions consistent among plants, Col‐0 and f6’h1 seedlings of similar size were equally transferred. This resulted in two plants of each genotype growing on each plate. Ten‐day‐old plants were inoculated along the primary root with 60–70 freshly hatched H. schachtii J2s each (Matera et al., 2021). The number of female and male nematodes was evaluated 12 days after inoculation using a Stereo Microscope (Leica, Germany), and mean values of nematode numbers were noted for the two plants on one plate. For scopolin quantification, the two plants on one same plate were pooled, for wild type (Col‐0) and f6’h1, all plants of one experiment were pooled for coumarin extraction.

Infection of soybean plants with F. virguliforme

Seeds of soybeans were germinated for 2 days on water‐soaked paper in the dark and then grown for 7 days on moist vermiculite (type Palabora, 3–6 mm; Isola Vermiculite GmbH). The roots of the soybean seedlings were then rinsed with water to remove substrate before placing three seedlings of the same genotype into one 50‐mL flacon tube containing 30 mL 0.5× Hoagland solution (see plant and fungal material) either supplemented with 30 million F. virguliforme spores or 0.5× Hoagland solution only, the latter of which served as controls. The plants were then grown in a growth cabinet for 7 days at 17 °C at a photoperiod of 16 h to facilitate infection. After another 7 days at 24 °C and 16 h photoperiod, the fresh weights of the aerial plant parts apart from cotyledons were determined.

The amount of fungal DNA in the root samples was quantified using a qRT‐PCR‐based method adapted from (Brouwer et al., 2003). Therefore, fungal DNA was extracted and set to 100 ng/μL and then diluted in a 4‐fold series with 100 ng/μL soybean DNA. A standard curve was prepared via qRT‐PCR using following primers FvTox1.F (5′‐GCAGGCCATGTTGGTTCTGTA‐3′) FvTox1.R (5′‐GCACGTAAAGTGAGTCGTCTCATC‐3′) (Wang et al., 2018). Fungal DNA content in infected plant material was calculated using the standard curve (Figure S12).

Mycotoxin‐induced ROS in soybean leaf discs and BY‐2 cell suspension culture

Mycotoxin‐induced ROS in soybean leaves was measured using a luminol‐based assay. Trifoliates of 19‐day‐old soybean plants were used for these experiments. Leaflets were excised, and leaf discs punched out using biopsy punches (5 mm, imtegra GmbH und Co. KG). Leaf discs were then placed in wells of a 96‐well‐plate filled with 200 μL ddH₂O and regenerated over night to avoid injury‐related stress responses. After 16 h, the water was replaced with 50 μL of fresh ddH₂O and rested for another 30 min. 50 μL of mastermix containing 2‐fold elicitor concentration, 40 μM L‐012 Luminol, 20 μg/mL HRP, and ddH₂O was added, and the plate directly transferred to the plate reader (BertholdTech CENTRO – MikroWin). Chitin served as a positive control and mastermix without elicitor as negative control. Measuring condition were as follows: 1 s/well, 90 s per cycle, mode: well after well, number of cycles: 27.

To indirectly quantify ROS accumulation in BY‐2 cells, a H₂DCFDA‐based assay was performed. Seven‐day‐old cell suspension cultures were adjusted to an equally packed cell volume and 1 mL of homogenous culture placed into a 24‐well plate. Elicitors were added to appropriate concentrations, and the cells incubated while shaking in the dark for 3 h (26 °C, 120 rpm). 200 μL of cell suspension was then transferred to a black NUNC96‐well plate (96F NUNCLEON DELTA lack microwell, ThermoFisher Scientific), and 2.5 μg/mL of H₂DCFDA was added. Following uptake into the plant cells, endogenous esterases cleave the non‐fluorescent H₂DCFDA, which is further oxidized to the fluorescent compound 2′‐7′‐dichlorofluorescin (Beyer et al., 2019). After incubating in the dark for another 30 min, this fluorescence was measured at 483–14 nm excitation and 530–30 nm emission in a CLARIOstar plate reader (BMG Labtech). Relative fluorescence units were set relative to the control to depict the ROS‐associated fluorescence.

Mycotoxin‐induced cell death in BY‐2 culture cells

For cell death visualization and quantification, cells were stained 4 h post‐treatment using propidium iodide (PI, 50 μg/mL, 200× stock in ddH₂O) and fluorescein diacetate (FDA, 25 μg/mL, 200× stock in acetone). Cells were analysed via confocal laser scanning microscopy (TCS SP Spectral Confocal Microscope, Leica Microsystems) shortly after staining cells for 10–15 min in reaction tubes on a rotating wheel. PI stains nuclei of dead cells, whereas FDA is converted by intracellular esterases resulting in the highly fluorescent fluorescein. The signal resulting from PI‐stained nuclei was detected at 561 nm excitation and 579–690 nm emission, and the fluorescein signal of living cells was detected at 488 nm excitation and 523–561 nm emission. Z‐stacks of several fields of view were recorded to ensure that cells in all layers were fully visible. To quantify the ratio between PI and FDA signal, the pixel sum in the respective channels was calculated using LAS X software (Leica Microsystems) and background‐corrected by subtracting the values from the control. Ratios calculated in this way indirectly represented viability of the cells. Higher values indicate a higher FDA signal compared to the PI signal, thereby showing higher viability of the cells.

Decompartmentalization of leaf cells

Fresh leaves were ground at room temperature and extracted at indicated time points using 1 mL of methanol (100%). Scopoletin and scopolin levels were determined using HPLC analysis, and the ratio subsequently calculated.

Conflict of interest

The authors H.S., U.C. and C.J.G.L. are inventors of project‐linked patent WO/2016124515. H.S. is an employee of BASF Plant Science Company GmbH. S.F.B. is an employee of BASF SE. U.C. and C.J.G.L. are employees of AgPrime GmbH.

Accession numbers

AtF6’H1 – AT3G13610.

Author contributions

A.B., S.F.B., S.B., C.J.G.L., U.C., H.S. and A.S.S. designed the experiments. A.B., S.F.B., V.W., S.L., S.B. and S.S.H. performed the experiments. A.B., C.J.G.L. and U.C. wrote the manuscript. J.G. and M.O. performed the GC–MS TOF analysis. All authors revsised the manuscript.

Supporting information

Figure S1 Scopoletin mainly accumulates in leaf veins and trichomes of AtF6’H1‐transgenic Arabidopsis plants. Transgenic Arabidopsis plants were exposed to UV light and photos taken to visualize scopoletin fluorescence.

Figure S2 Confirmation of the identity of scopoletin and scopolin in plant extracts by HPLC and GC–MS analysis. (a) Representative HPLC chromatograms (fluorescence at 365 nm excitation and 470 nm emission) of methanolic extracts from Arabidopsis and soybean wild‐type and AtF6‘H1‐overexpressing leaves and roots, authentic standards of esculin, scopolin, esculetin and scopoletin. (b) Verification of scopoletin and scopolin by GC–MS. Shown are the extracted spectra of the peaks of scopoletin and scopolin. Two numbers are given above each peak of the extracted spectra. The upper number indicates the m/z value, the lower represents the intensity.

Figure S3 Scopoletin and scopolin accumulate in seeds of AtF6‘H1‐overexpressing plants. Scopoletin and scopolin accumulate in AtF6’H1‐transgenic Arabidopsis (a) and soybean (b) seeds but not or only in low amounts in seeds of the respective wild‐type controls. Scopoletin and scopolin were quantified by HPLC. Shown are average values and SD from three seed extracts per transgenic event.

Figure S4 No phenotypic differences can be observed between wild‐type and coumarin‐accumulating soybean plants. Wild‐type and F6’H1‐overexpressing soybean plants were grown in long‐day conditions. Photos of six representative plants per genotype were taken 3.5 weeks after sowing. The white scale bar represents 10 cm.

Figure S5 Scopolin levels in roots of AtF6‘H1‐overexpressing Arabidopsis correlate with reduced number of male nematodes at the root. Shown is the average infection severity + SD of groups containing more (group >200) or less (group <200) than 200 μg/g FW scopolin in the root. Differences between groups >200 and <200 are statistically significant (t‐test; P < 0.05). Three independent experiments with at least four plates (two plants per plate) per genotype were performed.

Figure S6 AtF6‘H1‐overexpressing soybean lines show no obvious sudden death syndrome (SDS) tolerance of roots. Wild‐type plants and two independent AtF6‘H1‐overexpressing soybean events were hydroponically grown for 7 days. Plant roots were inoculated with F. virguliforme spores. Root biomass (fresh weight) of F. virguliforme‐infected plants was calculated relative to mock‐inoculated plants at 14 days after inoculation. Shown are min‐to‐max boxplots with all data points, average values (+) and SD of three independent experiments with nine plants per genotype and treatment. No significant differences to the wild type (control) were detected according to Dunnett's multiple comparisons test with P ≤ 0.05.

Figure S7 Scopoletin, in a dose‐dependent manner, reduces F. virguliforme biomass production in vitro. Growth medium was supplemented with 0.2% DMSO (control) or the indicated concentration of scopoletin in 0.2% DMSO, inoculated with 4 × 10⁵ F. virguliforme spores per mL. Mycelial dry weight was determined 7 days later. Shown are min‐to‐max boxplots with all data points, average values (+) and SD of three independent experiments. Significant differences to the negative control are marked with an asterisk (Students t‐test; P < 0.05).

Figure S8 Detached leaves of wild‐type and AtF6’H1‐overexpressing soybean plants exhibit no difference in water loss. Trifoliates from soybean wild‐type and transgenic scopoletin‐hyperaccumulating plants were detached and exposed to constant light at 26 °C. The relative weight was calculated relative to the fresh weight at time 0 min for each trifolium during 4 h. Mean ± SD shown from three experiments.

Figure S9 AtF6‘H1‐overexpression slightly but non‐significantly reduces SBR symptoms at late stages of infection. SBR symptoms were visually scored on three independent AtF6‘H1‐transgenic events of soybean genotype CD215 and azygous control plants at the indicated time points after planting. The experiment was performed in a test plot at Campinas, Brazi,l and consisted of three replicates/plots per event in a randomized design. Shown is the average infection severity of three replicates per event and time point. Differences were not significant according to statistical analysis with a mixed linear model in which the block was random, and test material (event and null) was fixed.

Figure S10 S. exigua larvae preferentially feed on wild type, rather than transgenic soybean leaf discs. Twenty‐seven leaf discs per indicated genotype were subjected to S. exigua feeding. (a) Representative photos of WT (wild type) and transgenic leaf discs 24 h of feeding. (b) The consumed leaf area was assessed and presented relative to the control. Shown are min‐to‐max boxplots with all data points, average values (+) and SD of three independent experiments. Asterisks indicate statistical difference between samples of WT and transgenic plants (t‐test, P ≤ 0.01).

Figure S11 HPLC gradient. Solvents used were ddH₂O and acetonitrile both supplemented with 1.5% acetic acid. The run time was 27 min at a flow rate of 0.8 mL/min.

Figure S12 Standard relationship of fungal DNA dilutions and C_T value in qRT‐PCR analysis. Genomic DNA of F. virguliforme was mixed with that of soybean and C_T values determined by qRT‐PCR.

Table S1 List of HPLC components used in the study. All components were from Shimadzu.

Table S2 List of primers used in the study. Blue colour indicates attB overhangs.