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. 2013 Oct 28;15(2):174–184. doi: 10.1111/mpp.12077

Arabidopsis GOLDEN2‐LIKE (GLK) transcription factors activate jasmonic acid (JA)‐dependent disease susceptibility to the biotrophic pathogen Hyaloperonospora arabidopsidis, as well as JA‐independent plant immunity against the necrotrophic pathogen Botrytis cinerea

Jhadeswar Murmu 1,3, Michael Wilton 2,4, Ghislaine Allard 1, Radhey Pandeya 1, Darrell Desveaux 2, Jas Singh 1, Rajagopal Subramaniam 1,✉
PMCID: PMC6638812  PMID: 24393452

Summary

Arabidopsis thaliana  GOLDEN2‐LIKE (GLK1 and 2) transcription factors regulate chloroplast development in a redundant manner. Overexpression of AtGLK1 (35S:AtGLK1) in Arabidopsis also confers resistance to the cereal pathogen Fusarium graminearum. To further elucidate the role of GLK transcription factors in plant defence, the Arabidopsis glk1 glk2 double‐mutant and 35S:AtGLK1 plants were challenged with the virulent oomycete pathogen Hyaloperonospora arabidopsidis (Hpa) Noco2. Compared with Col‐0, glk1 glk2 plants were highly resistant to Hpa  Noco2, whereas 35S:AtGLK1 plants showed enhanced susceptibility to this pathogen. Genetic studies suggested that AtGLK‐mediated plant defence to Hpa  Noco2 was partially dependent on salicylic acid (SA) accumulation, but independent of the SA signalling protein NONEXPRESSOR OF PATHOGENESIS‐RELATED 1 (NPR1). Pretreatment with jasmonic acid (JA) dramatically reversed Hpa  Noco2 resistance in the glk1 glk2 double mutant, but only marginally affected the 35S:AtGLK1 plants. In addition, overexpression of AtGLK1 in the JA signalling mutant coi1‐16 did not increase susceptibility to Hpa  Noco2. Together, our GLK gain‐of‐function and loss‐of‐function experiments suggest that GLK acts upstream of JA signalling in disease susceptibility to Hpa  Noco2. In contrast, glk1 glk2 plants were more susceptible to the necrotrophic fungal pathogen Botrytis cinerea, whereas 35S:AtGLK1 plants exhibited heightened resistance which could be maintained in the absence of JA signalling. Together, the data reveal that AtGLK1 is involved in JA‐dependent susceptibility to the biotrophic pathogen Hpa  Noco2 and in JA‐independent resistance to the necrotrophic pathogen B. cinerea.

Introduction

The GOLDEN2‐LIKE (GLK) genes are members of the GARP domain superfamily of transcription factors (TFs) (Riechmann et al., 2000). GLK was originally identified as G2 in maize (Hall et al., 1998) and was shown to be capable of transactivating the expression of a reporter gene in yeast (Rossini et al., 2001). Arabidopsis thaliana GLK1 (AtGLK1, GPRI1, At2g20570) and GLK2 (AtGLK2, GPRI2, At5g44190) (Fitter et al., 2002), also known as G‐BOX binding proline‐rich region interacting factors (GPRI) 1 and 2 (Tamai et al., 2002), are similar to type B ARABIDOPSIS RESPONSE REGULATORS (ARRs), which are families of plant proteins related to prokaryotic response regulators of two‐component systems (Riechmann et al., 2000). The AtGLK1 and AtGLK2 genes have been shown to act redundantly to regulate chloroplast development (Fitter et al., 2002; Yasumura et al., 2005). The loss‐of‐function glk1 glk2 double mutant is pale green in all photosynthetic tissues, with fewer thylakoid grana, whereas the loss of either AtGLK1 or AtGLK2 alone does not produce a pale green phenotype (Fitter et al., 2002). The AtGLK proteins also act in a cell autonomous manner to coordinate and maintain the photosynthetic apparatus within individual cells (Waters et al., 2008), and help to co‐regulate and synchronize the expression of nuclear photosynthetic genes to optimize photosynthetic capacity in varying environmental and developmental conditions (Waters et al., 2009). AtGLKs have also been shown to bind to promoters of several photosynthetic genes to upregulate the transcription of chloroplast genes (Waters et al., 2009). More recently, it has been observed that AtGLK2 can participate with LONG HYPOCOTYL (HY5) to achieve maximal effect in root greening in Arabidopsis (Kobayashi et al., 2012; Nakamura et al., 2009). In addition to binding to promoters, AtGLK1 also interacts with proline‐rich activation domains of G‐BOX binding bZIP factors, suggesting that GLKs may regulate other related plant processes (Tamai et al., 2002). Indeed, AtGLK1 overexpression has been shown to play a role in pathogen response (Savitch et al., 2007; Schreiber et al., 2011), in nitrogen use efficiency (Gutiérrez et al., 2008), in light‐ and brassinosteroid‐induced chloroplast development (Nakamura et al., 2009; Yu et al., 2011) and in root hormonal signalling (Kobayashi et al., 2012). Significantly, it has been observed recently that GLKs can interact with ANAC092 (ORE‐1) to modulate leaf senescence in Arabidopsis (Rauf et al., 2013).

Constitutive overexpression of AtGLK1 (35S:AtGLK1) confers enhanced resistance to Fusarium graminearum, a nonhost fungal pathogen in Arabidopsis (Savitch et al., 2007; Schreiber et al., 2011), suggesting that AtGLK1 may play a role in plant defence. In addition, an interactome network screening of plant pathogen effectors identified AtGLK1 to interact with the pathogen effector AvrRpm1‐like protein, although this interaction has not been confirmed directly (Mukhtar et al., 2012). The AtGLKs can also interact with ANAC092 to modulate senescence (Rauf et al., 2013), and with the involvement of ANAC092 in jasmonic acid (JA)/ethylene (ET) signalling (Bu et al., 2008) and in disease resistance (Al‐Daoud and Cameron, 2011). It is possible that GLKs may also be involved in responses to pathogen attack. To further elucidate the roles of AtGLK1 in plant pathogen responses, we used the ectopic overexpression of AtGLK1 (35S:AtGLK1) and the double knockout of AtGLK1 and AtGLK2 (glk1 glk2) to study the responses to the oomycete pathogen Hyaloperonospora arabidopsidis (Hpa) Noco2, an obligate biotroph causing downy mildew in the Arabidopsis host plant, and to Botrytis cinerea, a necrotrophic pathogen to Arabidopsis. Studies have indicated that the salicylic acid (SA) pathway plays a vital role in plant defence against biotrophic and hemibiotrophic pathogens, including Hpa (reviewed by Bari and Jones, 2009; Glazebrook, 2005). In addition, SA and JA defence pathways operating against different pathogens can be mutually antagonistic (Spoel et al., 2003; Van der Does et al., 2013; van Wees et al., 2003) or can operate in a synergistic fashion depending on the nature of the pathogen, thus underscoring the complexity of the signal transduction network involved in plant defence (Beckers and Spoel, 2006; Ferrari et al., 2003; Kunkel and Brooks, 2002; Mur et al., 2006; Schenk et al., 2000). Therefore, we introduced SA and JA signalling mutations into glk1 glk2 and 35:AtGLK1 plants to dissect the role of GLKs in plant defence. We showed that, compared with Col‐0, the glk1 glk2 double mutant exhibited elevated resistance to Hpa Noco2, whereas 35S:AtGLK1 was highly susceptible. Epistasis analyses with genes involved in the SA and JA signalling pathways suggested that the AtGLK1‐mediated defence to Hpa Noco2 was partially dependent on SA accumulation, but independent of NONEXPRESSOR OF PATHOGENESIS‐RELATED 1 (NPR1). The studies also indicated that AtGLK1 requires the JA receptor gene CORONATINE INSENSITIVE 1 (COI1) to promote disease susceptibility to Hpa Noco2. In contrast, 35S:AtGLK1 plants exhibited enhanced resistance to the necrotrophic fungal pathogen B. cinerea which was independent of COI1.

Results

Atglk1/glk2‐induced resistance to Hpa  Noco2 is partially dependent on SA accumulation, but independent of the SA signalling pathway

The glk1 glk2 double mutant and the overexpression of AtGLK1 (35S:AtGLK1) were evaluated for their response to Hpa Noco2 infection. Compared with Col‐0, the glk1 glk2 double‐mutant plants exhibited a strong restriction to Hpa Noco2 growth (Fig. 1a). Plants with mutations in either AtGLK1 (glk1) or AtGLK2 (glk2) displayed similar levels of Hpa Noco2 growth to the wild‐type Col‐0, suggesting that the resistance to Hpa Noco2 is the result of synergistic effects of both glk1 and glk2 (Fig. S1, see Supporting Information). This is consistent with observations of the redundant roles of AtGLK1 and AtGLK2 in chloroplast development (Fitter et al., 2002; Yasumura et al., 2005). Resistance to Hpa Noco2 in Arabidopsis is known to be mediated through the SA signalling pathway (Donofrio and Delaney, 2001; Lawton et al., 1995; Nawrath and Métraux, 1999; Sanchez et al., 2012), and this was verified here by enhanced susceptibility of Col‐0 plants expressing the bacterial enzyme salicylate hydroxylase (NahG), which catalyses SA to catechol (Fig. 1a). As the glk1 glk2 double‐mutant plants were highly resistant to Hpa Noco2, we were interested to determine whether this was the result of constitutive activation of the SA signalling pathway. First, we measured the levels of SA in uninfected plants. The results showed a dramatic 50% reduction in the glk1 glk2 double‐mutant plants compared with either the Col‐0 or 35S:AtGLK1 plants (Fig. 1b). After infection with Hpa Noco2, PATHOGENESIS‐RELATED 1 (PR1) (At2g14610) was significantly lower in the glk1 glk2 plants compared with either Col‐0 or the 35S:AtGLK1 plants (Fig. 1c). This suggests that SA signalling may not be a factor in the resistance to Hpa Noco2 in glk1 glk2 plants. These observations were verified by the expression of NahG in the glk1 glk2 double mutant (glk1 glk2 NahG) and by the introgression of the SA signalling mutant npr1‐1 into the glk1 glk2 double‐mutant plants (glk1 glk2 npr1‐1). As expected, NahG transgenic plants were susceptible to Hpa Noco2 (Fig. 1d) and the introduction of NahG into the glk1 glk2 double mutant (glk1 glk2 NahG) only partially rescued the Hpa Noco2 susceptibility of NahG plants (Fig 1d). Similarly, introgression of the SA signalling mutant npr1‐1 into the glk1 glk2 double‐mutant plants (glk1 glk2 npr1‐1) did not overcome the resistance observed in the glk1 glk2 double‐mutant plants (Fig. 1d). These observations confirmed that the resistance observed in the glk1 glk2 mutant plants was only partially the result of SA accumulation, and not mediated through NPR1. Correspondingly, in the 35S:AtGLK1 plants, genes involved in the SA signalling pathway, such as ENHANCED DISEASE SUSCEPTIBILITY 1 (EDS1), PHYTOALEXIN DEFICIENT4 (PAD4), ACCELERATED CELL DEATH6 (ACD6) and the SA biosynthesis enzyme ISOCHORISMATE SYNTHASE 1 (ICS1) accumulated to higher levels compared with those in Col‐0 plants (Fig. 2). This suggested that the susceptible phenotype observed in the 35S:AtGLK1 plants was not the result of impairments in SA biosynthesis or signalling.

Figure 1.

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Resistance to Hyaloperonospora arabidopsidis (Hpa) Noco2 is negatively regulated by AtGLK1 and AtGLK2 and is NPR1 independent. (a) One‐week‐old wild‐type (WT) (Col‐0), glk1 glk2 double‐mutant, 35S:AtGLK1 and NahG plants were inoculated with 1 × 105/mL Hpa  Noco2 spores. At 7 days post‐infection (dpi), the spores were quantified from three samples of 25–30 seedlings per sample. The data from one representative experiment are shown and the error bars represent the standard error (SE). Similar trends were observed in three independent biological replicates. (b) Quantification of salicylic acid (SA), jasmonic acid‐isoleucine (JA‐Ile) and JA in 10‐day‐old seedlings of the WT (Col‐0), glk1 glk2 double mutant and 35S:AtGLK1. Analyses were performed on three separate biological samples and error bars represent SE. * and ** indicate significant differences between the WT and mutants at P < 0.01 and P < 0.05, respectively (Student's t‐test). (c) PATHOGENESIS‐RELATED 1 (PR1) transcript levels relative to UBIQUITIN 5 (UBQ5) were measured by quantitative reverse transcription‐polymerase chain reaction (qRT‐PCR) from RNA isolated at 72 h post‐infection (hpi) with Hpa  Noco2. Similar trends in relative transcript levels were observed in two independent biological replicates. (d) Analyses show that the resistance to Hpa  Noco2 in the glk1 glk2 double mutant is partially dependent on SA accumulation, but independent of NONEXPRESSOR OF PATHOGENESIS‐RELATED 1 (NPR1). The experimental details are similar to (a), but included npr1‐1, glk1 glk2 npr1‐1 and glk1 glk2 NahG. FW, fresh weight.

Figure 2.

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Transcript analyses of salicylic acid (SA) pathway genes on Hyaloperonospora arabidopsidis (Hpa) Noco2 infection. Quantitative reverse transcription‐polymerase chain reaction (qRT‐PCR) of the indicated gene transcripts relative to UBIQUITIN 5 (UBQ5) as internal standard at 72 h post‐infection (hpi) with Hpa  Noco2. (a) ENHANCED DISEASE SUSCEPTIBILITY 1 (EDS1) transcripts; (b) PHYTOALEXIN DEFICIENT4 (PAD4) transcripts; (c) ISOCHORISMATE SYNTHASE 1 (ICS1) transcripts; (d) ACCELERATED CELL DEATH6 (ACD6) transcripts. The data are representative of one experiment. Similar trends in relative transcript levels were obtained from three independent biological replicates. WT, wild‐type.

Pretreatment with SA partially rescues 35S:AtGLK1 susceptibility to Hpa  Noco2

Pretreatment with SA has been used to induce resistance against biotrophs, including Hpa Noco2 (Kunkel and Brooks, 2002; Ryals et al., 1996). Accordingly, pretreatment with SA conferred increased resistance to Hpa Noco2 in Col‐0 plants, whereas npr1‐1 plants, which are defective in SA signalling, did not respond to SA treatment (Fig. 3a). A similar pretreatment of 35S:AtGLK1 plants with SA only marginally reduced the susceptibility to Hpa Noco2 (Fig. 3a). Interestingly, pretreatment also reduced resistance in glk1 glk2 plants, but these plants were still considerably more resistant than untreated Col‐0 plants. Together, these results suggested that the activation of the SA pathway led to partial rescue of the susceptible phenotype observed in the 35S:AtGLK1 plants. The pretreatment, however, did not compromise the ability to express PR1 in any of the mutant lines, and the expression was comparable with that of Col‐0 treated with SA (Fig. 3b). This suggested that the SA signalling pathway was functional in these mutants, but was not recruited to provide resistance to Hpa Noco2 (Fig. 3a).

Figure 3.

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Pretreatment with salicylic acid (SA) only marginally affects Hyaloperonospora arabidopsidis (Hpa) Noco2 susceptibility in 35S:AtGLK1 plants. (a) One‐week‐old seedlings were misted with either sterile water + 0.01% Silwet‐77, which served as a control (−SA), or 1 mm SA in water + 0.01% Silwet‐77 (+SA). The seedlings were covered with a clear plastic dome for 24 h prior to spray inoculation with 1 × 105/mL Hpa  Noco2 spores. At 7 days post‐infection (dpi), spores were quantified from three samples with 25–30 seedlings per sample. Data from a representative experiment are shown and the error bars represent the standard error (SE). Similar trends were observed in three independent biological replicates. FW, fresh weight. (b) RNA was isolated from the samples from (a) and quantitative reverse transcription‐polymerase chain reaction (qRT‐PCR) was performed relative to UBIQUITIN 5 (UBQ5). Similar trends in relative transcript levels were obtained from three independent biological replicates. PR1, PATHOGENESIS‐RELATED 1; WT, wild‐type.

Arabidopsis susceptibility to Hpa  Noco2 requires both JA signalling and AtGLK1

Resistance to Hpa Noco2 is known to be achieved either by the activation of the SA pathway or, alternatively, by suppression of the JA pathway (Argueso et al., 2012; Clarke et al., 1998; Lawton et al., 1995; Li et al., 2001; Massoud et al., 2012; Murray et al., 2002; Rairdan and Delaney, 2002). In the absence of evidence to support SA‐mediated resistance in the glk1 glk2 plants, we hypothesized that the JA signalling pathway may be inoperative or compromised in the glk1 glk2 plants or, alternatively, the JA pathway is constitutively active in the 35S:AtGLK1 plants. To test this hypothesis, we pretreated both glk1 glk2 and 35S:AtGLK1 plants with methyl jasmonate (MeJA) and observed for disease phenotypes on Hpa Noco2 infection. Pretreatments with 0.1 mm MeJA dramatically reversed the resistance to Hpa Noco2 in the glk1 glk2 mutant plants (Fig. 4). The pretreatment, however, did not increase significantly the already susceptible 35S:AtGLK1 plants (Fig. 4). To obtain further insights into the role of GLKs in the JA signalling pathway, we crossed coi1‐16, a mutant that does not bind JA or JA‐isoleucine (Ile), into the glk1 glk2 double‐mutant and 35S:AtGLK1 plants. The coi1‐16 mutant was similar to Col‐0 plants in susceptibility and, as expected, did not respond to JA application to increase this susceptibility (Fig. 4). However, the introduction of glk1 glk2 into the coi1‐16 mutant plants (coi1‐16 glk1 glk2) completely restored the resistance to Hpa Noco2 to at least the level of the glk1 glk2 mutant (Fig. 4). This suggested that the activation of the JA pathway by MeJA treatment was necessary for susceptibility and was mitigated by the presence of GLKs. This was further supported by the observation that plants overexpressing AtGLK1, which were susceptible with or without MeJA pretreatment, were completely resistant in the coi1‐16 mutant background (35S:AtGLK1 coi1‐16), and indicated that AtGLK1 and JA/JA‐Ile perception through COI1 were both necessary for susceptibility to Hpa Noco2 (Fig. 4). Measurements of JA and JA‐Ile showed no significant differences between the Col‐0 and 35S:AtGLK1 plants, and further supported the notion that the integration of GLKs and the JA signalling pathway was required to observe a susceptible phenotype against the Hpa Noco2 pathogen (Fig. 1b).

Figure 4.

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Susceptibility to Hyaloperonospora arabidopsidis (Hpa) Noco2 is dependent on jasmonic acid (JA) signalling. One‐week‐old seedlings were misted with either sterile water + 0.01% Silwet‐77, which served as a control (–JA), or 0.1 mm methyl jasmonate in water + 0.01% Silwet‐77 (+JA). The seedlings were covered with a clear plastic dome for 24 h prior to spray inoculation with 1 × 105/mL Hpa  Noco2 spores. At 7 days post‐infection (dpi), spores were quantified from three samples with 25–30 seedlings per sample. Data from one representative experiment are shown and the error bars represent the standard error (SE). Similar trends were observed in three independent biological replicates. FW, fresh weight; WT, wild‐type.

AtGLK1 confers resistance to B. cinerea independent of JA signalling

To corroborate our findings that AtGLK1, in addition to JA signalling, may be an integral component in plant defence, we employed another pathosystem that requires JA signalling for defence. Unlike, Hpa Noco2, resistance to the necrotroph B. cinerea in Arabidopsis is known to be mediated through the JA signalling pathway (Glazebrook, 2005; Grant and Jones, 2009). To determine whether genetic manipulation of GLKs can alter responses to this pathogen, excised leaves from the glk1 glk2 double‐mutant and 35S:AtGLK1 plants were infected with B. cinerea. We observed that 35S:AtGLK1 plants were considerably more resistant than Col‐0 plants to B. cinerea and that the resistance was maintained even after 5 days (Fig. 5a–c). In contrast, glk1 glk2 plants were significantly more susceptible than Col‐0 at 48 h after infection and the lesion continued to grow even at 5 days post‐infection (dpi), similar to the coi1‐16 control plants (Fig. 5a–c). The coi1‐16 glk1 glk2 triple mutant was equally susceptible to the individual coi1‐16 and glk1 glk2 mutants (Fig. 5a–c). In contrast, overexpression of AtGLK1 in the coi1‐16 mutant background (coi1‐16 35S:AtGLK1) resulted in a highly resistant phenotype similar to 35S:AtGLK1 plants (Fig. 5a–c). Furthermore, this resistance to B. cinerea was not accompanied by changes in transcript levels of JA signalling markers, such as VEGETATIVE STORAGE PROTEIN 2 (VSP2), Thionin 2.1 (Thi2.1), PLANT DEFENSIN 1 and 2 (PDF1 and PDF2) (compare coi1‐16 35S:AtGLK1 infected with coi1‐16 infected in Table S3, see Supporting Information). This suggested that, unlike the Arabidopsis response to the biotroph Hpa Noco2, overexpression of AtGLK1 obviated the requirement of COI1 in resistance to the necrotroph B. cinerea. Conceivably, AtGLK1 can function to facilitate resistance to B. cinerea either downstream or independent of COI1.

Figure 5.

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AtGLK1 positively regulates resistance to Botrytis cinerea. Detached leaves from 4‐week‐old plants were inoculated with 6 μL of Botrytis cinerea spores (5 × 105 spores/mL) and photographed at the indicated times. Lesion sizes were measured and quantified by ImageJ software (http://www.nih.gov/ij). Data represent the means ± standard deviation (SD) from 30 infected leaves. (a) 48 h post‐infection (hpi); (b) 5 days post‐infection (dpi); (c) lesion measurements at 48 hpi and 5 dpi. The data are representative of three independent biological experiments. Scale bar, 1 cm in (a) and (b). WT, wild‐type.

Discussion

Previously, we have reported that constitutive overexpression of AtGLK1 confers enhanced resistance to the necrotrophic fungal pathogen F. graminearum in Arabidopsis (Savitch et al., 2007; Schreiber et al., 2011). The present study aimed to further elucidate the role of AtGLK1 in disease resistance and to gain insights into plant responses to biotrophic and necrotrophic pathogens. SA biosynthesis and signalling have been demonstrated to be critical for resistance against plant pathogens (Argueso et al., 2012; Bowling et al., 1994; Clarke et al., 1998, 2000; Delaney et al., 1994; Glazebrook et al., 1996; Li et al., 2001; Murray et al., 2002; Rairdan et al., 2001; Vlot et al., 2009; Wildermuth et al., 2001). In addition, SA signalling has been identified to positively regulate resistance against Hpa (Lawton et al., 1994; Sanchez et al., 2012). Here, our observation that glk1 glk2 double‐mutant plants are highly resistant to Hpa Noco2, whereas 35S:AtGLK1 plants are highly susceptible, when compared with Col‐0 (Fig. 1a), suggests that AtGLK1 is a negative regulator of resistance to Hpa Noco2. We have provided evidence that the AtGLK1‐regulated Arabidopsis response to Hpa Noco2 is partially dependent on SA accumulation, but independent of SA signalling. Further, we observed that the introduction of NahG into the glk1 glk2 double mutant (glk1 glk2 NahG) only partially rescued the Hpa Noco2 susceptibility of NahG (Fig. 1d). The results with NahG, however, should be interpreted with caution as the expression of this gene has been shown to exhibit pleiotropic effects (Heck et al., 2003; van Wees and Glazebrook, 2003). As the glk1 glk2 npr1‐1 triple mutant can rescue npr1‐1 susceptibility to Hpa Noco2 (Fig. 1d), we suggest that the regulation of Hpa Noco2 resistance in the glk1 glk2 double mutant is independent of SA signalling. In conjunction with the observation that the expression of PR1, EDS1, PAD4 and ICS1 genes indicates activation of the SA pathway in the highly susceptible 35S:AtGLK1 plants (Fig. 2), the results suggest that AtGLK1‐mediated defence responses to Hpa Noco2 are predominantly SA independent.

JA signalling and AtGLK1 enhance susceptibility to Hpa  Noco2 in Arabidopsis

We have demonstrated that either overexpression of AtGLK1 or activation of the JA pathway by MeJA in the glk1 glk2 double‐mutant plants leads to disease outcome (Fig. 4). Furthermore, SA pretreatment only partially rescues the disease phenotype in the AtGLK1 plants (Fig. 3a). Together, these results suggest that overexpression of GLK1 may antagonize SA‐induced resistance against Hpa Noco2. When these results are juxtaposed with other observations, such as the resistant phenotype in coi1 35S:AtGLK1 plants, it reinforces the notion that GLK may participate in JA–SA antagonism. JA–SA antagonism in the pathogen response is well established (Robert‐Seilaniantz et al., 2011; Thaler et al., 2012).

AtGLK1 is a positive regulator of defence against B. cinerea which is independent of JA signalling

Plants overexpressing AtGLK1 (35S:AtGLK1) were highly resistant to B. cinerea. Conversely, glk1 glk2 plants were highly susceptible (Fig. 5). It is generally recognized that resistance to biotrophs is mediated through SA signalling, whereas resistance to necrotrophs is mediated through JA signalling (Glazebrook, 2005). Plants impaired in JA signalling are more susceptible to necrotrophic fungal pathogens, including B. cinerea (Ferrari et al., 2007; La Camera et al., 2011; Rowe and Kliebenstein, 2008). As expected, coi1‐16 plants were highly susceptible to B. cinerea (Fig. 5). However, plants overexpressing AtGLK1 in the coi1‐16 background (coi1‐16 35S:AtGLK1) were highly resistant, suggesting that AtGLK1‐induced resistance is independent of COI1. This COI1 independence is further supported by observations of the lack of induced transcripts of JA signalling marker genes on infection with B. cinerea (Table S3; compare coi1‐16 and coi1‐16 35S:AtGLK1). COI1‐independent processes have been observed to be involved in resistance to necrotrophs, especially against B. cinerea (Birkenbihl et al., 2012; Ferrari et al., 2007; Laluk et al., 2011; Pré et al., 2008; Tang et al., 2007) and soil‐borne fungal pathogens (Ralhan et al., 2012; Thatcher et al., 2009). Coincidentally, 35S:AtGLK1 plants showed attenuated constitutive levels of VSP2, Thi2.1 and PDF transcripts compared with Col‐0 and glk1 glk2 (Table S3). The association of attenuated levels of JA signalling marker transcripts with enhanced resistance to B. cinerea has been reported for the Atmyc2.2 mutant and the anac019 anac055 double mutant (Anderson et al., 2004; Bu et al., 2008). Recently, AtGLKs have been reported to interact directly with ANAC092 (ORE‐1) to modulate chloroplast degradation in senescence (Rauf et al., 2013). It is therefore conceivable that AtGLK1 can act independently and downstream of COI1 by the suppression of JA signalling through interactions with AtMYC2, ANAC019 and/or ANAC055, and thus play a critical role in plant defence.

A proposed model for AtGLK1 in plant defence

The combined results of Hpa Noco2 resistance in the absence of AtGLKs or the JA‐receptor COI1 and its reversibility by MeJA application suggest that GLK1 functions upstream of JA signalling (Fig. 6). In the absence of GLKs (glk1 glk2) and on infection with Hpa Noco2, the JA pathway is compromised, resulting in resistance (possibly by the removal of JA antagonism to SA). On infection with B. cinerea, resistance is provided by the presence of GLK1 in an as yet unknown mechanism which is independent of JA signalling. The requirement of GLK for resistance is evidenced by the susceptibility of glk1 glk2 to B. cinerea, and the independence from JA signalling is demonstrated in the recovery of resistance in the coi1‐16 mutant by GLK1 overexpression (coi1‐16 35S:AtGLK1) and by the absence of the expression of target genes of JA signalling (Table S3). The mechanism of how AtGLKs influence plant defences is not known, but we could envisage a scenario in which ‘effectors’ from Hpa Noco2 and B. cinerea could target GLKs, leading to JA‐dependent susceptibility against the biotroph and JA‐independent resistance against the necrotroph (Fig. 6). AtGLK1 has been reported previously to interact with AvrRpm1, an effector from the bacterial pathogen (Mukhtar et al., 2012).

Figure 6.

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A proposed model for the role of AtGLK1 in plant defence. AtGLK may be a potential target for virulence factors from Hyaloperonospora arabidopsidis (Hpa) Noco2 and Botrytis cinerea. If GOLDEN2‐LIKE (GLK) acts as a negative regulator of the jasmonic acid (JA) pathway, targeting it will activate this pathway and promote susceptibility to Hpa  Noco2 and resistance against the biotroph. The experimental evidence suggests that the promotion of susceptibility to Hpa  Noco2 requires the intact JA signalling pathway (open arrows). In contrast, this pathway is not required for resistance to B. cinerea (filled arrows). COI1, CORONATINE INSENSITIVE 1.

Experimental Procedures

Plant materials and growth conditions

Arabidopsis thaliana (Col‐0) single‐knockout lines of glk1 (At2g20570) and glk2 (At5g44190) and the glk1 glk2 double‐knockout line, N9805, N9806, N9807, respectively, were obtained from the Nottingham Arabidopsis Stock Centre (NASC), Nottingham, UK. The Col‐0 plants overexpressing GLK1 (35S:AtGLK1) were constructed as described previously (Savitch et al., 2007). Arabidopsis thaliana (Col‐0), glk1, glk2, glk1 glk2, npr1‐1, glk1 glk2 npr1‐1, glk1 glk2 NahG, NahG, coi1‐16, coi1‐16 glk1 glk2, coi1‐16 35S:AtGLK1 and 35S:AtGLK1 seeds were surface sterilized and subjected to vernalization at 4 °C for 2 days. The seeds were planted in ProMix soil and grown in 16 h light and 8 h dark at 21 °C at 60% relative humidity and under a light intensity of 150 μmol photons/m2/s. Plants were fertilized every second week. Double and triple mutants were generated by hand crossing, and segregating F2 populations were polymerase chain reaction (PCR) genotyped to identify the homozygous double and triple mutants. The details are described in Figs S2–S5 (see Supporting Information).

PCR genotyping

PCR genotyping primers are listed in Table S1 (see Supporting Information) and the results of PCR genotyping of the mutants are shown in Figs S2–S5.

Pathogen infection, microscopy, photographs and imaging

Spores of Hpa strain Noco2 were freshly harvested from Col‐0 seedlings in sterile water and adjusted to 105/mL for all the infection experiments. One‐week‐old Arabidopsis seedlings were spray inoculated with Hpa Noco2 using a spray gun (Preval, Coal City, IL, USA), and the plants were covered with a clear plastic dome to maintain high humidity (>95%) and grown at 16 °C in 9 h light/15 h dark for 7 days in an Adaptis A1000 growth chamber (Conviron, Winnipeg, MB, Canada). At 7 dpi, Hpa Noco2 spores from infected plants were harvested in sterile water and quantified. For gene expression analyses, infected plants were harvested at 72 h post‐infection (hpi) and quick frozen in liquid N2 for RNA isolation. Botrytis cinerea strain B191 spores were harvested from potato dextrose agar (PDA) plates in sterile water, filtered through four layers of sterile cheesecloth and washed twice in sterile water. The spores were resuspended in potato dextrose broth (PDB) and diluted to a concentration of 5 × 105/mL for pathogenicity tests. Detached leaves from 4‐week‐old plants were placed on three layers of moist sterile Whatman filter paper (Fisher Scientific, ON, Canada) inside a sterile plastic Petri dish. Leaves were inoculated individually with 6 μL of B. cinerea spores at the centre of the leaf and the Petri dish was sealed with surgical tape to maintain humidity. Plates were incubated in the dark for the first 12 h and then transferred to a growth chamber set at 21 °C and 16 h light/8 h dark. The diseased leaves were photographed at varying times to observe lesion spread. Lesion sizes were measured from the digital photographs using ImageJ software (http://www.nih.gov).

Quantitative reverse transcription‐polymerase chain reaction (qRT‐PCR)

Total RNA was isolated by Trizol reagent (Sigma‐Aldrich, Oakville, ON, Canada). Two micrograms of RNA were converted to cDNA by High Capacity cDNA Reverse Transcription Kits (Applied Biosystems). All of the qRT‐PCRs were performed in triplicate by an Applied Biosystems Power SYBR Green Kit and the Applied Biosystems StepOne Plus Real‐Time PCR System according to the manufacturer's instructions (Applied Biosystems, Burlington, ON, Canada). For relative quantification, a standard curve for each primer set was created and UBIQUITIN 5 (UBQ5) was used as internal control for standardization between samples. Data were analysed by StepOne 2.1 software. A list of the genes and primers used in qRT‐PCR is given in Table S2 (see Supporting Information). Analyses of transcript level changes after B. cinerea infection (Table S3) were performed as follows: 4‐week‐old soil‐grown plants were spray inoculated with either PDB (mock) or B. cinerea spores (2.5 × 105/mL) in PDB; the plants were covered with a clear plastic dome to maintain high humidity and incubated in the dark for the first 12 h, and later transferred to a growth chamber at 21 °C with a 16‐h light/8‐h dark photoperiod. Leaf samples were harvested at 48 hpi and quick frozen in liquid N2 for RNA isolation. Gene transcripts were analysed by qRT‐PCR relative to UBQ5 as internal standard. P < 0.05 was considered to be statistically significant.

Hormone analyses

Leaves (approximately 500 mg) from 10‐day‐old Col‐0, glk1 glk2 and 35S:AtGLK1 plants were harvested and immediately frozen in liquid nitrogen for SA, JA and JA‐IIe analyses. The frozen fresh plant materials were extracted with methanol–water–glacial acetic acid (3 mL, 90:9:1, v/v/v) to which the internal standards were added (100 mL of solution containing acetonitrile–water 50:50, v/v, with 0.1% formic acid, containing 1 ng/mL of 3,4,5,6‐d4‐2‐hydroxybenzoic acid and 0.5 ng/mL of 2,2‐d2‐JA). Following sonication (5 min) and incubation in an orbital shaker (4 °C, 5 min), the samples were centrifuged at 266 300 g for 10 min to pellet the debris. The supernatants were transferred to clean tubes and the pellets were resuspended in the extraction solution (2 mL), and the procedure was repeated. The supernatants from individual samples were combined with the initial extracted volume and the pellets were resuspended in methanol (1 mL). The extraction step was repeated a third time. After the supernatants had been combined, methanol was evaporated under reduced pressure. On ice, aqueous NaOH was added to each sample to obtain a basic pH (1 mL, 0.3 M), and was then further extracted with dichloromethane (3 mL). The aqueous layers were transferred to a clean tube, and the organic layer was re‐extracted with aqueous NaOH (2 mL). On ice, the combined aqueous layers were acidified with 5% aqueous HCl (1 mL) and then extracted with a mixture of ethyl acetate and cyclohexane (1 mL, 1:1, v/v). The organic phases were collected and the aqueous phases were extracted a second time with the same mixture (0.5 mL). The organic fractions were pooled and the solvent was evaporated under a constant nitrogen stream. Prior to mass spectrometric analysis, the samples were reconstituted in a mixture of methanol and water (200 mL, 30:70, v/v) containing 0.1% formic acid, to which external standards were added (100 ng of 1,2,3,4,5,6‐13C6‐2‐hydroxybenzoic acid and 50 ng of 12,12,12‐d3‐JA). SA and JA were purchased from Sigma‐Aldrich; JA‐Ile was purchased from OlChemIm Ltd. (Olomouc, Czech Republic). The preparation of standards and analyses were carried out at the National Research Council of Canada, Saskatoon, Canada. Deuterated forms of the hormones which were used as internal standards, 2,2‐d2‐JA and 12,12,12‐d3‐JA, were prepared as described by Galka et al. (2005). The preparation of 1,2,3,4,5,6‐13C6‐2‐hydroxybenzoic acid has been presented elsewhere (P. W. S. Galka et al., unpublished data, National Research Council, Sask, Canada). Calibration curves were created for all compounds of interest. In the absence of an appropriate deuterium‐ or 13C‐labelled internal standard, JA‐Ile was quantified using a calibration curve built on its response against d3‐JA. Quality control samples (QCs) were run alongside the tissue samples. For the SA/JA analysis, the analytical ACQUITY UPLC® HSS C18 column (2.1 mm × 100 mm, 1.8 μm; Waters Limited, ON, Canada) was used. The compounds were eluted from the column with a mixture of solvents comprising 1% formic acid in high‐performance liquid chromatography (HPLC)‐grade water (mobile phase A) and 1% formic acid in HPLC‐grade methanol (mobile phase B), using a gradient mode. Analyses were performed by ultra‐performance liquid chromatography‐electrospray tandem mass spectrometry (UPLC/ESI‐MS/MS) using a Waters ACQUITY UPLC system, equipped with a binary solvent delivery manager and a sample manager coupled to a Waters Micromass Quattro Premier XE quadrupole tandem mass spectrometer via a Z‐spray interface. MassLynx™ and QuanLynx™ (Micromass, Manchester, UK) were used for data acquisition and analyses.

Supporting information

Fig. S1  AtGLK1 and AtGLK2 redundantly regulate the Hyaloperonospora arabidopsidis (Hpa) Noco2 resistance in Arabidopsis. One‐week‐old seedlings were inoculated with 1 × 105/mL Hpa Noco2 spores. At 7 days post‐infection (dpi), the spores were quantified from three samples of 25–30 seedlings per sample. The data from one representative experiment are shown and the error bars represent the standard error (SE). Similar trends were observed in three independent biological replicates.

Fig. S2 Polymerase chain reaction (PCR) genotyping of the glk1 glk2 NahG plant. (A) The top gel shows the presence of the dSpm insertion in the glk1.2 mutation detected with the 2bgs2 + Spm5 primer pair at 58 °C after 35 cycles of PCR, and the bottom gel shows the presence of the wild‐type (WT) allele for AtGLK1 detected with the GLK1‐Promoter‐F1 + AtGLK1‐R primer pair at 58 °C after 35 cycles of PCR to confirm the homozygous status of the glk1.2 mutation. The asterisk (*) represents nonspecific PCR products. (B) The top gel shows the presence of the dSpm insertion in the glk2.1 mutation detected with the Spm1 + AtGLK2‐R primer pair at 62 °C + gradient primer annealing after 35 cycles of PCR, and the bottom gel shows the presence of the WT allele for AtGLK2 detected with the AtGLK2‐F + AtGLK2‐R primer pair at 62 °C + gradient primer annealing with 35 cycles of PCR to confirm the homozygous status of the glk2.1 mutation. PCR genotyping of glk1.2 and glk2.1 mutations was performed according to Fitter et al. (2002). (C) The presence of the NahG transgene was confirmed with the NahG‐F + NahG‐R primer pair at 58 °C after 35 cycles of PCR generating a 700‐bp product. Aliquots of 5 μL of PCR products from each genotype were separated on a 2% (w/v) agarose gel.

Fig. S3 Polymerase chain reaction (PCR) genotyping of the glk1 glk2 npr1‐1 triple mutant. (A) The top gel shows the presence of the dSpm insertion in glk1.2 detected with the 2bgs2 + Spm5 primer pair at 58 °C after 35 cycles of PCR, and the bottom gel shows the presence of the wild‐type (WT) allele for AtGLK1 detected with the GLK1‐Promoter‐F1 + AtGLK1‐R primer pair at 58 °C after 35 cycles of PCR to confirm the homozygous status of the glk1.2 mutation. The asterisk (*) indicates nonspecific PCR products. (B) The top gel shows the presence of the dSpm insertion in glk2.1 detected with the Spm1 + AtGLK2‐R primer pair at 62 °C + gradient primer annealing with 35 cycles of PCR, and the bottom gel shows the presence of the WT allele for AtGLK2 detected with the AtGLK2‐F + AtGLK2‐R primer pair at 62 °C + gradient primer annealing after 35 cycles of PCR to confirm the homozygous status of the glk2.1 mutation. Aliquots of 5 μL of PCR product from each genotype were separated on a 2% (w/v) agarose gel. PCR genotyping of glk1.2 and glk2.1 mutations was performed according to Fitter et al. (2002). (C) NPR1 was amplified using the npr1‐1 F + npr1‐1R dCAPs primer pair at 58 °C and generated a 252‐bp product, which was digested with NlaIII (NEB) and separated on a 3% agarose gel. The PCR product was digested in the WT of the NPR1 allele, but not in the npr1‐1 mutant allele.

Fig. S4 Polymerase chain reaction (PCR) genotyping of the coi1‐16 glk1 glk2 triple mutant. (A) The top gel shows the presence of the dSpm insertion in the glk1.2 mutation detected with the 2bgs2 + Spm5 primer pair at 58 °C primer annealing with 35 cycles of PCR, and the bottom gel represents the presence of the wild‐type (WT) allele for GLK1 detected with the GLK1‐Promoter‐F1 + AtGLK1‐R primer pair at 58 °C primer annealing with 35 cycles of PCR to confirm the homozygous status of the glk1.2 mutation. The asterisk (*) represents nonspecific PCR products. (B) The top gel shows the presence of the dSpm insertion in the glk2.1 mutation detected with the Spm1 + AtGLK2‐R primer pair at 62 °C + gradient primer annealing with 35 cycles of PCR, and the bottom gel shows the presence of the WT allele for GLK2 detected with the AtGLK2‐F + AtGLK2‐R primer pair at 62 °C + gradient primer annealing with 35 cycles of PCR to confirm the homozygous status of the glk2.1 mutation. PCR genotyping of the glk1.2 and glk2.1 mutation was performed according to Fitter et al. (2002). (C) COI1 in the WT was detected using the coi1‐16 P2 + coi1‐16 P4 primer pair at 64 °C primer annealing with 26 cycles of PCR. As coi1‐16 is a point mutation, the WT allele for COI1 also generated a PCR product with the coi1‐16 P2 + coi1‐16 P3 primer pair; however, the coi1‐16 P2 + coi1‐16 P4 primer pair generated a PCR product only in the WT allele and not in the coi1‐16 mutant allele. PCR genotyping of the coi1‐16 mutant was performed according to Adams and Turner (2010). Aliquots of 5 μL of PCR products from each genotype were separated on a 2% (w/v) agarose gel.

Fig. S5 Polymerase chain reaction (PCR) genotyping of the coi1‐16 35S:AtGLK1 plant. (A) Presence of the genomic GLK1 and cDNA of the inserted AtGLK1 transgene. The GLK1 genomic fragment was detected using the AtGLK1‐F + GLK1 genomic‐R primer pair at 58 °C after 35 cycles of PCR. (B) Detection of the coi1‐16 mutation using the coi1‐16 P2 + coi1‐16 P3 primer pair at 64 °C after 26 cycles of PCR. Aliquots of 5 mL of PCR products from each genotype were separated on a 2% (w/v) agarose gel. (C) COI1 detected in the wild‐type (WT) using the coi1‐16 P2 + coi1‐16 P4 primer pair at 64 °C after 26 cycles of PCR. Aliquots of 5 mL of PCR products from each genotype were separated on a 2% (w/v) agarose gel. As coi1‐16 is a point mutation, the WT allele for COI1 also generated a PCR product with the coi1‐16 P2 + coi1‐16 P3 primer pair. However, the coi1‐16 P2 + coi1‐16 P4 primer pair generated PCR products only in the WT, but not in the coi1‐16 mutant alleles. PCR genotyping of the coi1‐16 mutant was performed according to Adams and Turner (2010).

Table S1 Primer combinations used to genotype mutant and transgenic plants.

Table S2 List of quantitative reverse transcription‐polymerase chain reaction (qRT‐PCR) primers.

Table S3 Transcript levels of VEGETATIVE STORAGE PROTEIN 2 (VSP2), Thionin 2.1 (Thi2.1), PLANT DEFENSIN 1 and 2 (PDF1 and PDF2) after infection with Botrytis cinerea.

Click here for additional data file. (631.8KB, zip)

Acknowledgements

This project was supported in part by the Agriculture and Agri‐Food Canada Crop Genomics Initiative. The authors wish to thank Dr Irina Zaharia of the Plant Biotechnology Institute, National Research Council, Saskatoon, SK, Canada for hormone analyses.

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Associated Data

This section collects any data citations, data availability statements, or supplementary materials included in this article.

Supplementary Materials

Fig. S1  AtGLK1 and AtGLK2 redundantly regulate the Hyaloperonospora arabidopsidis (Hpa) Noco2 resistance in Arabidopsis. One‐week‐old seedlings were inoculated with 1 × 105/mL Hpa Noco2 spores. At 7 days post‐infection (dpi), the spores were quantified from three samples of 25–30 seedlings per sample. The data from one representative experiment are shown and the error bars represent the standard error (SE). Similar trends were observed in three independent biological replicates.

Fig. S2 Polymerase chain reaction (PCR) genotyping of the glk1 glk2 NahG plant. (A) The top gel shows the presence of the dSpm insertion in the glk1.2 mutation detected with the 2bgs2 + Spm5 primer pair at 58 °C after 35 cycles of PCR, and the bottom gel shows the presence of the wild‐type (WT) allele for AtGLK1 detected with the GLK1‐Promoter‐F1 + AtGLK1‐R primer pair at 58 °C after 35 cycles of PCR to confirm the homozygous status of the glk1.2 mutation. The asterisk (*) represents nonspecific PCR products. (B) The top gel shows the presence of the dSpm insertion in the glk2.1 mutation detected with the Spm1 + AtGLK2‐R primer pair at 62 °C + gradient primer annealing after 35 cycles of PCR, and the bottom gel shows the presence of the WT allele for AtGLK2 detected with the AtGLK2‐F + AtGLK2‐R primer pair at 62 °C + gradient primer annealing with 35 cycles of PCR to confirm the homozygous status of the glk2.1 mutation. PCR genotyping of glk1.2 and glk2.1 mutations was performed according to Fitter et al. (2002). (C) The presence of the NahG transgene was confirmed with the NahG‐F + NahG‐R primer pair at 58 °C after 35 cycles of PCR generating a 700‐bp product. Aliquots of 5 μL of PCR products from each genotype were separated on a 2% (w/v) agarose gel.

Fig. S3 Polymerase chain reaction (PCR) genotyping of the glk1 glk2 npr1‐1 triple mutant. (A) The top gel shows the presence of the dSpm insertion in glk1.2 detected with the 2bgs2 + Spm5 primer pair at 58 °C after 35 cycles of PCR, and the bottom gel shows the presence of the wild‐type (WT) allele for AtGLK1 detected with the GLK1‐Promoter‐F1 + AtGLK1‐R primer pair at 58 °C after 35 cycles of PCR to confirm the homozygous status of the glk1.2 mutation. The asterisk (*) indicates nonspecific PCR products. (B) The top gel shows the presence of the dSpm insertion in glk2.1 detected with the Spm1 + AtGLK2‐R primer pair at 62 °C + gradient primer annealing with 35 cycles of PCR, and the bottom gel shows the presence of the WT allele for AtGLK2 detected with the AtGLK2‐F + AtGLK2‐R primer pair at 62 °C + gradient primer annealing after 35 cycles of PCR to confirm the homozygous status of the glk2.1 mutation. Aliquots of 5 μL of PCR product from each genotype were separated on a 2% (w/v) agarose gel. PCR genotyping of glk1.2 and glk2.1 mutations was performed according to Fitter et al. (2002). (C) NPR1 was amplified using the npr1‐1 F + npr1‐1R dCAPs primer pair at 58 °C and generated a 252‐bp product, which was digested with NlaIII (NEB) and separated on a 3% agarose gel. The PCR product was digested in the WT of the NPR1 allele, but not in the npr1‐1 mutant allele.

Fig. S4 Polymerase chain reaction (PCR) genotyping of the coi1‐16 glk1 glk2 triple mutant. (A) The top gel shows the presence of the dSpm insertion in the glk1.2 mutation detected with the 2bgs2 + Spm5 primer pair at 58 °C primer annealing with 35 cycles of PCR, and the bottom gel represents the presence of the wild‐type (WT) allele for GLK1 detected with the GLK1‐Promoter‐F1 + AtGLK1‐R primer pair at 58 °C primer annealing with 35 cycles of PCR to confirm the homozygous status of the glk1.2 mutation. The asterisk (*) represents nonspecific PCR products. (B) The top gel shows the presence of the dSpm insertion in the glk2.1 mutation detected with the Spm1 + AtGLK2‐R primer pair at 62 °C + gradient primer annealing with 35 cycles of PCR, and the bottom gel shows the presence of the WT allele for GLK2 detected with the AtGLK2‐F + AtGLK2‐R primer pair at 62 °C + gradient primer annealing with 35 cycles of PCR to confirm the homozygous status of the glk2.1 mutation. PCR genotyping of the glk1.2 and glk2.1 mutation was performed according to Fitter et al. (2002). (C) COI1 in the WT was detected using the coi1‐16 P2 + coi1‐16 P4 primer pair at 64 °C primer annealing with 26 cycles of PCR. As coi1‐16 is a point mutation, the WT allele for COI1 also generated a PCR product with the coi1‐16 P2 + coi1‐16 P3 primer pair; however, the coi1‐16 P2 + coi1‐16 P4 primer pair generated a PCR product only in the WT allele and not in the coi1‐16 mutant allele. PCR genotyping of the coi1‐16 mutant was performed according to Adams and Turner (2010). Aliquots of 5 μL of PCR products from each genotype were separated on a 2% (w/v) agarose gel.

Fig. S5 Polymerase chain reaction (PCR) genotyping of the coi1‐16 35S:AtGLK1 plant. (A) Presence of the genomic GLK1 and cDNA of the inserted AtGLK1 transgene. The GLK1 genomic fragment was detected using the AtGLK1‐F + GLK1 genomic‐R primer pair at 58 °C after 35 cycles of PCR. (B) Detection of the coi1‐16 mutation using the coi1‐16 P2 + coi1‐16 P3 primer pair at 64 °C after 26 cycles of PCR. Aliquots of 5 mL of PCR products from each genotype were separated on a 2% (w/v) agarose gel. (C) COI1 detected in the wild‐type (WT) using the coi1‐16 P2 + coi1‐16 P4 primer pair at 64 °C after 26 cycles of PCR. Aliquots of 5 mL of PCR products from each genotype were separated on a 2% (w/v) agarose gel. As coi1‐16 is a point mutation, the WT allele for COI1 also generated a PCR product with the coi1‐16 P2 + coi1‐16 P3 primer pair. However, the coi1‐16 P2 + coi1‐16 P4 primer pair generated PCR products only in the WT, but not in the coi1‐16 mutant alleles. PCR genotyping of the coi1‐16 mutant was performed according to Adams and Turner (2010).

Table S1 Primer combinations used to genotype mutant and transgenic plants.

Table S2 List of quantitative reverse transcription‐polymerase chain reaction (qRT‐PCR) primers.

Table S3 Transcript levels of VEGETATIVE STORAGE PROTEIN 2 (VSP2), Thionin 2.1 (Thi2.1), PLANT DEFENSIN 1 and 2 (PDF1 and PDF2) after infection with Botrytis cinerea.

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