E3 ligase AtAIRP5/GARU regulates drought stress response by stimulating SERINE CARBOXYPEPTIDASE-LIKE1 turnover

Abstract

Ubiquitination is a major mechanism of eukaryotic posttranslational protein turnover that has been implicated in abscisic acid (ABA)-mediated drought stress response. Here, we isolated T-DNA insertion mutant lines in which ABA-insensitive RING protein 5 (AtAIRP5) was suppressed, resulting in hyposensitive ABA-mediated germination compared to wild-type Arabidopsis (Arabidopsis thaliana) plants. A homology search revealed that AtAIRP5 is identical to gibberellin (GA) receptor RING E3 ubiquitin (Ub) ligase (GARU), which downregulates GA signaling by degrading the GA receptor GID1, and thus AtAIRP5 was renamed AtAIRP5/GARU. The atairp5/garu knockout progeny were impaired in ABA-dependent stomatal closure and were markedly more susceptible to drought stress than wild-type plants, indicating a positive role for AtAIRP5/GARU in the ABA-mediated drought stress response. Yeast two-hybrid, pull-down, target ubiquitination, and in vitro and in planta degradation assays identified serine carboxypeptidase-like1 (AtSCPL1), which belongs to the clade 1A AtSCPL family, as a ubiquitinated target protein of AtAIRP5/GARU. atscpl1 single and atairp5/garu-1 atscpl1-2 double mutant plants were more tolerant to drought stress than wild-type plants in an ABA-dependent manner, suggesting that AtSCPL1 is genetically downstream of AtAIRP5/GARU. After drought treatment, the endogenous ABA levels in atscpl1 and atairp5/garu-1 atscpl1-2 mutant leaves were higher than those in wild-type and atairp5/garu leaves. Overall, our results suggest that AtAIRP5/GARU RING E3 Ub ligase functions as a positive regulator of the ABA-mediated drought response by promoting the degradation of AtSCPL1.

An Arabidopsis RING E3 ubiquitin ligase positively regulates abscisic acid-mediated drought stress response by promoting 26S proteasome-dependent turnover of a putative acyltransferase.

Introduction

Land plants are inherently confronted with adverse environmental stresses, such as extreme temperatures, heavy metals, high salinity, and water deficits. These abiotic stresses can seriously affect plant development and growth, which leads to decreased crop yields (Lobell and Field, 2007; Fedoroff et al., 2010; Zhu, 2016). To cope with these hostile conditions, land plants have developed sophisticated defense mechanisms (Takahashi et al., 2020). Abscisic acid (ABA) is a key phytohormone involved in abiotic stress responses by regulating diverse subsets of defense genes (Finkelstein, 2013; Cotelle and Leonhardt, 2019; Todaka et al., 2019). Because environmental stress conditions often fluctuate rapidly, the ABA response must be strictly regulated to cope with these dynamic unfavorable growth conditions.

The ubiquitin (Ub)-26S proteasome system (UPS) is a major mechanism of eukaryotic posttranslational protein turnover. The UPS regulates crucial cellular processes, including cell cycle progression, hormone responses, and defense responses against biotic and abiotic stresses (Stone, 2014; Yu et al., 2016). Target proteins that are designated for degradation through the UPS are commonly modified by the attachment of a poly-Ub chain. The ubiquitination pathway proceeds in three consecutive steps that are, respectively, catalyzed by E1 Ub-activating enzymes, E2 Ub-conjugating enzymes, and E3 Ub ligases (Smalle and Viestra, 2004; Lee and Kim, 2011). Because substrate specificity is primarily determined by E3 Ub ligases, these enzymes have attracted much interest (Xu and Xue, 2019; Tal et al., 2020).

Previous reports have indicated that Really Interesting New Gene (RING) domain-containing E3 Ub ligases participate in the ABA-mediated drought stress response (Lyzenga and Stone, 2012; Shu and Yang, 2017). Arabidopsis (Arabidopsis thaliana) ABA-insensitive RING protein 1 (AtAIRP1), which is a cytosolic- and nuclear-localized RING-type E3 ligase, positively regulates the ABA-dependent drought stress response by ubiquitinating the importin-β protein AtKPNB1 (Ryu et al., 2010; Oh et al., 2020). A loss-of-function mutation in salt- and drought-induced RING finger1 (SDIR1) E3 Ub ligase resulted in a phenotype of drought stress susceptibility (Zhang et al., 2007). SDIR1 and AtAIRP2 ubiquitinate and promote the degradation of the transcription activator/pterin dehydratase ATP1/SDIRIP1 in the ABA and salt stress responses (Cho et al., 2011; Zhang et al., 2015; Oh et al., 2017). AtAIRP3/LOG2 plays a positive role in the response to drought stress by downregulating the cysteine protease RD21 and the ABA coreceptor ABI1 (Kim and Kim, 2013; Pan et al., 2020). The cytosolic E3 Ub ligase AtAIRP4 is a positive factor in ABA-mediated drought avoidance and is a negative factor in the salinity tolerance response (Yang et al., 2016). The E3 ligases RGLG1 and RGLG5 induce turnover of phosphatase PP2CA, a key repressor of ABA signaling, and thus positively regulate the ABA response (Wu et al., 2016). In contrast, the RING E3 Ub ligases DRIP1 and RGLG2 act as negative factors in the response to drought stress by enhancing proteasomal degradation of the drought-responsive transcription factors DREB2A and ERF53, respectively (Qin et al., 2008; Cheng et al., 2012). MIEL1 ubiquitinates the transcription factor MYB96 and negatively regulates ABA signaling (Lee and Seo, 2016). These findings suggest that RING E3 Ub ligases play either a positive or negative role in the ABA and drought stress responses by inducing UPS-mediated turnover of their target proteins.

In this study, we identified T-DNA insertion loss-of-function mutant lines in the At2g40830 locus, which showed ABA hyposensitivity during the germination stage. The predicted At2g40830 protein possessed a single typical RING motif in its middle region and was named AtAIRP5. Unexpectedly, AtAIRP5 was found to be identical to GA receptor RING E3 Ub ligase (GARU), which prevents gibberellin (GA) signaling by degrading the GA receptor GID1 (Nemoto et al., 2017). Thus, AtAIRP5 was renamed AtAIRP5/GARU. Nemoto et al. (2017) reported that the expression levels of AtAIRP5/GARU increased by water and salt stress in seeds. They further showed that AtAIRP5/GARU modulates sensitivity of GA response in seeds under the salt stress. The atairp5/garu knockout lines displayed impaired ABA-dependent stomatal movement, which resulted in a phenotype highly susceptible to drought stress as compared to wild-type Arabidopsis plants. Our results indicated that AtAIRP5/GARU targets serine carboxypeptidase-like1 (AtSCPL1) protein for the UPS-mediated degradation. Contrary to the atairp5/garu mutants, the atscpl1 loss-of-function mutant plants were more tolerant to drought stress in an ABA-dependent manner than wild-type plants. The atairp5/garu-1 atscpl1-2 double mutant progeny also showed greater tolerance to drought than wild-type plants, indicating that AtSCPL1 is downstream of AtAIRP5/GARU. Suppression of AtAIRP5/GARU and AtSCPL1 resulted in the decrease and increase in the endogenous ABA amounts, respectively, in response to drought stress, suggesting that the opposite phenotypes of the atairp5/garu and atscpl1 mutants to drought stress were due, at least in part, to a difference in endogenous ABA levels. In conclusion, our data indicate that AtAIRP5/GARU functions as a positive regulator of the ABA-mediated drought stress response by downregulating AtSCPL1. These results are discussed considering the suggestion that the RING E3 Ub ligase AtAIRP5/GARU plays a negative role in GA signaling and a positive role in ABA signaling in Arabidopsis.

Results

Identification of the ABA- and drought-induced AtAIRP5/GARU RING-type E3 Ub ligase in Arabidopsis

Previous germination assays revealed that T-DNA insertion atairp1, atairp2, and atairp3/log2 knockout Arabidopsis mutants in which these RING E3 Ub ligase genes were inactivated were hyposensitive to ABA when compared to wild-type plants at the germination stage (Ryu et al., 2010; Cho et al., 2011; Kim and Kim, 2013). Figure 1A shows that a loss-of-function mutation in the At2g40830 locus resulted in ABA-insensitive germination, and the phenotype of these plants was similar to those of atairp1 and atairp2 plants. The At2g40830 gene (GenBank accession No. NM_201922) comprised 2,293 bp with a single intron, was located on chromosome 2, and was named AtAIRP5 (Figure 1B). We identified three different T-DNA insertion loss-of-function mutant alleles of AtAIRP5, and the T-DNA insertions were mapped to the coding region (atairp5-1; SALK_037121) and the 5′-untranslated region (atairp5-2; SALK_127503 and atairp5-3; SALK_137684) of the AtAIRP5 gene (Figure 1B). Homozygous atairp5 progeny were verified by genotyping PCR (Supplemental Figure S1A). Reverse transcription (RT)–PCR revealed that the atairp5-1 and atairp5-3 alleles were null mutants, whereas a low level of AtAIRP5 transcript was detected in the atairp5-2 plants (Supplemental Figure S1B). Thus, the atairp5-1 and atairp5-3 null mutant lines were used for subsequent phenotypic analyses.

Figure 1.

Identification and characterization of drought-induced AtAIRP5/GARU RING-type E3 Ub ligase. A, Isolation of an ABA-insensitive RING E3 Ub ligase mutant (at2g40830) at the germination stage. Wild-type and T-DNA-inserted mutant plants, in which putative RING-type E3 Ub ligase genes were suppressed, were grown on MS medium containing 0 or 0.25-μM ABA. The germination percentage was calculated based on cotyledon greening at 10 days after sowing. The atairp1 and atairp2 mutant plants served as positive controls for the ABA-hyposensitive phenotype during the germination stage. Scale bars = 0.5 cm. B, Schematic representation of AtAIRP5/GARU and the three atairp5/garu T-DNA insertion mutant alleles (atairp5/garu-1; SALK_037121, atairp5/garu-2; SALK_127503, and atairp5/garu-3; SALK_137684). The T-DNA insertion sites are marked with inverted triangular boxes. The arrows represent the position and direction of the primers used for genotyping PCR and RT–PCR analyses. Light gray bars, dark gray bars, and solid lines represent the 5′- and 3′-untranslated regions, coding region, and intron, respectively, of the AtAIRP5/GARU gene. The C3H2C3-type RING motif is indicated in the middle of the predicted AtAIRP5/GARU protein. C, RT-qPCR analysis of AtAIRP5/GARU expression in response to ABA, high salinity, and water stress. Light-grown, 8-day-old wild-type seedlings were subjected to 100-μM ABA (2 h), 300-mM NaCl (2 h), and dehydration (4 h). RAB18 was used as a positive control for ABA treatment. RD29A was used as a positive control for drought and salt stress. The transcript levels of each gene were normalized to that of glyceraldehyde-3-phosphate dehydrogenase C subunit. Error bars are the sd of three biologically independent experiments (n > 25); Double asterisks indicate statistical significance [^**P < 0.01, by one-way ANOVA with post hoc Tukey’s honestly significant difference (HSD) test]. D, ABA-mediated germination assay of wild-type, atairp5/garu-1, and atairp5/garu-3 plants. Sterilized seeds of each genotype were germinated with different concentrations (0, 0.6, 0.9, and 1.2 μM) of ABA. Scale bars = 0.5 cm. E, Quantitative analysis of the germination data shown in (D). Error bars are the sd of three biologically independent experiments (n > 25); Double asterisks indicate statistical significance (^**P < 0.01, by one-way ANOVA with post hoc Tukey’s HSD test).

The deduced AtAIRP5 protein contained a single typical RING motif in its middle region (Figure 1B; Supplemental Figure S2). A database search of The Arabidopsis Information Resource (http://www.arabidopsis.org) revealed that AtAIRP5 was identical to GARU (Nemoto et al., 2017). Thus, AtAIRP5 was renamed AtAIRP5/GARU. Nemoto et al. (2017) reported that AtAIRP5/GARU functions as a negative regulator of GA signaling by degrading the GA-receptor GID1 via the UPS in seeds under salt stress. In wild-type seeds, approximately two-fold increase in the expression of AtAIRP5/GARU was detected in response to water and salt stress (Nemoto et al., 2017). RT–qPCR analysis showed that, in light-grown 8-day-old seedlings, AtAIRP5/GARU transcript levels were enhanced by ABA (5.6-fold), drought (14.1-fold), and NaCl (16.3-fold) treatments (Figure 1C). AtAIRP5/GARU has been previously shown to be an active Ub ligase (Nemoto et al., 2017). Our in vitro self-ubiquitination assay also demonstrated that the full-length MBP-AtAIRP5/GARU fusion protein had E3 ligase activity in the presence of Ub, ATP, E1 (Arabidopsis UBA1), and E2 (Arabidopsis UBC8), as detected by both anti-MBP and anti-Ub antibodies (Supplemental Figure S3A). In contrast, a single amino acid substitution derivative (MBP-AtAIRP5/GARU^C208S), in which the conserved Cys-208 residue in the RING domain was replaced with Ser-208, did not show self-ubiquitination activity. A subcellular localization assay using AtAIRP5/GARU^C208S-green fluorescent protein (GFP) showed that AtAIRP5/GARU was predominantly present in the cytosol and nucleus of Nicotiana benthamiana leaf cells (Supplemental Figure S3B).

Because the atairp5/garu mutant lines were initially identified by their ABA-insensitive germination phenotype (Figure 1A), we repeated the seed germination assay using wild-type and atairp5/garu seeds with different concentrations (0, 0.6, 0.9, and 1.2 μM) of ABA. Germination ratios were determined as the percentages of cotyledon greening after 10 days of stratification. The results showed that wild-type and mutant (atairp5/garu-1 and atairp5/garu-3) seeds fully germinated in the absence of ABA. In the presence of 0.6 μM ABA, ∼95%, 97%, and 98% of the wild-type, atairp5/garu-1, and atairp5/garu-3 seeds germinated, respectively (Figure 1, D and E). The differences between the wild-type and mutant seeds became more pronounced as the ABA concentration increased, and in the presence of 0.9 μM ABA, ∼54%, 78%, and 76% of the wild-type, atairp5/garu-1, and atairp5/garu-3 seeds germinated, respectively. In the presence of 1.2-μM ABA, only 13% of the wild-type seeds germinated, whereas >25% of the atairp5/garu mutants germinated (Figure 1, D and E). These results revealed that suppression of AtAIRP5/GARU resulted in ABA insensitivity during the germination stage.

To further examine the role of AtAIRP5/GARU in the ABA response, we constructed 2×Flag-AtAIRP5/GARU-overexpressing transgenic Arabidopsis plants under the control of the cauliflower mosaic virus 35S promoter. Ectopic expression of the AtAIRP5/GARU transcript and protein was confirmed in two independent T3 transgenic plants (lines #1 and #2) by RT–PCR and immunoblot analysis, respectively (Supplemental Figure S4, A and B). In contrast to the atairp5/garu mutant plants, the AtAIRP5/GARU-overexpressing seedlings did not show a detectable phenotypic difference compared to the wild-type seedlings regarding ABA-mediated seed germination (Supplemental Figure S4, C and D). To examine whether 2×Flag-AtAIRP5/GARU was functional in planta, the 2×Flag-AtAIRP5/GARU construct was ectopically expressed in the atairp5/garu mutant plants (Supplemental Figure S5, A–C). The results of seed germination assays showed that the ABA-insensitive phenotype of atairp5/garu seedlings were efficiently rescued by constitutive expression of 2×Flag-AtAIRP5/GARU, which indicated that 2×Flag-AtAIRP5/GARU was functional in planta (Supplemental Figure S5, D and E). In contrast, 2×Flag-AtAIRP5/GARU^C208S was unable to complement the ABA-insensitive germination of atairp5/garu.

Loss-of-function mutants of AtAIRP5/GARU showed impaired ABA-mediated stomatal closure and hypersensitivity to drought stress

Expression of AtAIRP5/GARU was increased by ABA and drought treatments (Figure 1C), and the atairp5/garu mutant plants were hyposensitive to ABA during the germination stage (Figure 1, D and E). Therefore, we speculated that AtAIRP5/GARU is involved in the ABA-dependent drought stress response. To test this possibility, phenotypic assays were performed. First, we measured the stomatal movements of wild-type and atairp5/garu and atairp3/log2-2 mutant leaves. atairp3/log2-2 was used as a positive control for ABA insensitivity (Kim and Kim, 2013). The abaxial parts of the leaves were peeled off from light-grown 4-week-old mature plants, incubated with stomata opening solution (Kwak et al., 2003) for 4 h, and then treated with different concentrations of ABA (0, 1.0, and 10.0 μM) for 2 h. Without ABA, the average stomatal apertures (width to length ratio) of wild-type and mutant leaves were indistinguishable (Figure 2, A and B). As ABA concentration increased, wild-type leaves exhibited typical ABA-dependent stomatal closure, with average apertures of 0.137 ± 0.005 with 1.0-μM ABA and 0.113 ± 0.004 with 10.0-μM ABA. In contrast, the ABA-mediated stomatal movements of the atairp5/garu and atairp3/log2-2 mutant leaves were markedly reduced compared to those of wild-type plants in response to the same range of ABA concentrations. The average stomatal apertures of the atairp5/garu-1, atairp5/garu-3, and atairp3/log2-2 leaves were 0.166 ± 0.005, 0.171 ± 0.007, and 0.169 ± 0.007 with 1.0-μM ABA and 0.154 ± 0.007, 0.158 ± 0.005, and 0.165 ± 0.004 with 10.0-μM ABA, respectively (Figure 2, A and B).

Figure 2.

Phenotypic analyses of atairp5/garu loss-of-function mutant plants in response to ABA and dehydration stress. A, ABA-mediated stomatal closure of wild-type, atairp5/garu-1, atairp5/garu-3, and atairp3/log2-2 plants. Light-grown, 4-week-old rosette leaves of each genotype were incubated with ABA (0, 1.0, and 10.0 μM) for 2 h. atairp3/log2-2 was used as a positive control for ABA-hyposensitive stomatal closure. Stomatal apertures were observed using a light microscope. Scale bars = 10 μm. B, Quantification of the stomatal aperture data shown in (A). Stomatal apertures were measured as the ratio of the width to length using the ImageJ software. Error bars are the sd of three biologically independent experiments (n > 35); asterisks indicate statistical significance [^**P < 0.01, by one-way analysis of variance (ANOVA) with post hoc Tukey’s HSD test]. C, Drought sensitivity of atairp5/garu-1 and atairp5/garu-3 mutant plants compared with wild-type plants. Light-grown, 18-day-old mature plants of each genotype were exposed to dehydration stress for 11 days. Then, the survival of drought stress-treated plants was assessed after 3 days of re-watering. The images were digitally extracted for comparison. Scale bars = 2 cm. D, Leaf water loss assay. The aerial parts of wild-type, atairp5/garu-1, atairp5/garu-3, and atairp3/log2-2 plants were detached and incubated at room temperature for the indicated time points, and water loss rates were calculated as the percentage of initial FW of the aerial parts. Error bars are the sd of three biologically independent experiments (n > 12).

Next, wild-type and atairp5/garu mutant plants were subjected to dehydration stress. These plants were grown for 18 days in pots under normal growth conditions, and then irrigation was stopped for 11 days. After 3 days of re-watering, the survival ratios were assessed. Under our experimental conditions, 56.6% (34 of 60) of the wild-type plants resumed growth after irrigation, while only 15.0% (9 of 60) and 3.3% (2 of 60) of the atairp5/garu-1 and atairp5/garu-3 plants, respectively, survived (Figure 2C). These results suggest that malfunction of AtAIRP5/GARU in Arabidopsis markedly led to a reduced tolerance to water deficit. The aerial parts of the wild-type, atairp5/garu, and atairp3/log2-2 plants were detached and placed on open-lid petri dishes at room temperature. Then, the time-dependent reduction in the fresh weights (FWs) of the detached aerial parts was monitored over time (0–360 min). After 60 min of incubation, the wild-type, atairp5/garu, and atairp3/log2-2 plants retained ∼84.1%, 80.5%–81.0%, and 80.5% of their starting weights, respectively (Figure 2D). After 360 min of incubation, the wild-type, atairp5/garu, and atairp3/log2-2 plants were reduced to ∼61.9%, 56.7%–56.8%, and 56.1% of their starting weights, respectively. Thus, the aerial parts of the mutant plants lost water more rapidly than those of the wild-type plants. Taken together, the results of the phenotypic analyses in Figure 2 suggest that AtAIRP5/GARU plays a positive role in the ABA-mediated drought stress response.

AtAIRP5/GARU interacts with AtSCPL1

To unravel the molecular mechanism underlying the positive role of AtAIRP5/GARU in the ABA-mediated drought stress response, we performed a yeast two-hybrid screen using AtAIRP5/GARU as bait. The full-length AtAIRP5/GARU cDNA and a cDNA library of 3-day-old etiolated seedlings as prey were co-transformed into yeast strain AH109 and cultured in triple-minus selection medium (SD/–Trp/–Leu/–His). p53 (murine p53^72-390) + T-antigen and lambda (human lamin C^66–230) + T-antigen were used as positive and negative controls, respectively. We identified several positive yeast cells that grew in the selection medium, one of which contained a partial cDNA encoding the C-terminal region (amino acids 209–441) of AtSCPL1 (At5g36180) (Figure 3, A and B). Full-length AtSCPL1 possesses 441 amino acids (molecular mass, 50.0 kDa) with a putative 29 amino acid-long N-terminal signal peptide and a conserved catalytic triad comprising Ser, Asp, and His residues (Figure 3B). When a 35S:AtSCPL1-GFP fusion construct was transiently expressed in protoplasts prepared from Arabidopsis leaf cells using a polyethylene glycol (PEG)-based method, the fluorescent signal exhibited a web-like network pattern throughout the cytosol (upper part in Figure 3C). In addition, the cytosolic network pattern of the AtSCPL1-GFP signal closely overlapped with that of BiP-mRFP-HDEL, an ER marker protein, in N. benthamiana leaf epidermal cells (lower part in Figure 3C), indicating that AtSCPL1 is predominantly localized in the ER. The results of RT–qPCR analysis revealed that AtSCPL1 was downregulated by ABA, NaCl, and dehydration treatments (Figure 3D).

Figure 3.

Identification of AtSCPL1 as a putative interacting partner of AtAIRP5/GARU. A, Yeast two-hybrid screen using AtAIRP5/GARU as bait and a cDNA library prepared from 3-day-old etiolated seedlings as prey. Yeast cells (AH109) were plated on SD/–Leu/–Trp/–His medium and incubated at 30°C for 4 days. p53 + T-antigen and lambda + T-antigen were used as the positive and negative controls, respectively. B, Schematic representation of the predicted AtSCPL1 protein, which comprises 441 amino acids. The putative signal peptide (amino acids 1–29) and conserved catalytic triad (Ser-184, Asp-366, and His-419) are marked. A dashed line indicates a partial cDNA encoding the C-terminal region (amino acids 209–441) of AtSCPL1 that was initially identified by yeast two-hybrid screening. C, Subcellular localization assays of AtSCPL1 in Arabidopsis protoplasts and N. benthamiana leaf epidermal cells. Upper part, The AtSCPL1-GFP fusion protein was transiently expressed in Arabidopsis protoplasts prepared from wild-type leaf cells using a PEG-based method. Lower, The 35S:AtSCPL1-GFP fusion construct was introduced into N. benthamiana leaf epidermal cells along with 35S:BiP1-mRFP-HDEL (an ER marker) using the Agrobacterium-mediated infiltration method. The fluorescent signals were observed by confocal microscopy under dark-field or light-field conditions. Scale bars = 20 μm. D, RT–qPCR analysis of AtSCPL1 expression in response to ABA, high salinity, and water stress. Light-grown, 8-day-old wild-type seedlings were incubated in liquid MS medium containing 100-μM ABA (2 h) or 300-mM NaCl (2 h). For the water stress treatment, 8-day-old seedlings were exposed to dehydration stress (4 h) by opening the lid of the Petri dishes. The transcript levels of each gene were normalized to that of glyceraldehyde-3-phosphate dehydrogenase C subunit. Error bars are the sd of three biologically independent experiments (n > 25); asterisks indicate statistical significance (*P < 0.05 and **P < 0.01, by one-way ANOVA with post hoc Tukey’s HSD test).

The Arabidopsis AtSCPL family consists of 51 proteins with homology to SCPs belonging to the peptidase S10 family (Mugford and Milkowski, 2012). Despite their homology to carboxypeptidases, AtSCPLs do not possess peptidase activity (Milkowski and Strack, 2004; Stehle et al., 2009). Instead, the AtSCPL family exhibits acyltransferase activity (Lehfeldt et al., 2000; Sherley et al., 2001). These AtSCPL acyltransferase activities are involved in a broad spectrum of cellular processes, including defense responses to biotic and abiotic stresses, secondary metabolite pathways, membrane lipid metabolism, and conjugation of phytohormone (Liu et al., 2008; Mugford et al., 2009; Ciarkowska et al., 2019; Chen et al., 2020). AtSCPL1 belongs to clade IA of the AtSCPL family (Mugford and Milkowski, 2012; Supplemental Figures S6 and S7). Although several clade 1A AtSCPLs are known to be involved in sinapic acid metabolism in Arabidopsis, the biological role of AtSCPL1 has not yet been evaluated.

To verify and further characterize the interaction between AtAIRP5/GARU and AtSCPL1, we used two different AtSCPL1 derivatives: full-length AtSCPL1 and a signal peptide-deleted ΔSP-AtSCPL1 mature form. The results of a yeast two-hybrid assay showed that AtAIRP5/GARU failed to interact with full-length AtSCPL1 but could interact with ΔSP-AtSCPL1 in triple-minus selection medium (Figure 4A), suggesting that full-length AtSCPL1 might not be correctly processed to the mature form in yeast cells. Next, we conducted an in vitro pull-down assay. Bacterially expressed MBP, MBP-AtAIRP5/GARU, and 6×His-3×Myc-ΔSP-AtSCPL1 recombinant proteins were co-incubated with an anti-His affinity gel matrix. The resin-bound proteins were subjected to immunoblot analysis using anti-Myc and anti-MBP antibodies. As shown in Figure 4B, MBP-AtAIRP5/GARU, but not MBP, was co-immunoprecipitated with 6×His-3×Myc-ΔSP-AtSCPL1, indicating a physical interaction between AtAIRP5/GARU and ΔSP-AtSCPL1. The full-length 6×His-3×Myc-AtSCPL1 protein was precipitated in Escherichia coli cells, possibly due to its eukaryotic signal peptide; thus, it was excluded in the pull-down assay. We also performed an in vivo co-immunoprecipitation (co-IP) assay to examine the interaction in planta. 35S:AtSCPL1-GFP was infiltrated with or without 35S:2×Flag-AtAIRP5/GARU into N. benthamiana leaves using the Agrobacterium-mediated transformation method. Leaf crude extracts (containing 500 µg of total protein) were immunoprecipitated using a magnetic bead-bound anti-Flag antibody. The results revealed that AtSCPL1-GFP was precipitated by the anti-Flag antibody in the leaf extracts that expressed 2×Flag-AtAIRP5/GARU (left part in Figure 4C). In contrast, 2×Flag-AtAIRP5/GARU did not interact with GFP, which was used as a negative control (right part in Figure 4C). Thus, unlike in yeast cells, the full-length AtSCPL1 appeared to be correctly processed and interacted with AtAIRP5/GARU in planta.

Figure 4.

In vitro and in planta interactions between AtAIRP5/GARU and AtSCPL1. A, Yeast two-hybrid assay of AtAIRP5/GARU and AtSCPL1. The AtAIRP5/GARU-pGBKT7 + AtSCPL1-pGADT7 and AtAIRP5/GARU-pGBKT7 + ΔSP-AtSCPL1-pGADT7 constructs were co-introduced into yeast AH109 cells. p53 + T-antigen was used as a positive control, and lambda + T-antigen was used as a negative control. B, In vitro pull-down assay. Bacterially expressed MBP, MBP-AtAIRP5/GARU, and 6×His-3×Myc-ΔSP-AtSCPL1 recombinant proteins were purified and incubated with Ni-NTA agarose resin. The resin-bound proteins were eluted, separated by sodium dodecyl sulfate-polyacrylamide gel electrophoresis (SDS-PAGE), and detected with anti-Myc and anti-MBP antibodies. C, In vivo co-IP assay. The 35S:AtSCPL1-GFP (left) and 35S:GFP (right) constructs were transiently expressed with or without 35S:2×Flag-AtAIRP5/GARU in N. benthamiana epidermal cells. The crude extracts were incubated with magnetic bead-bound anti-Flag antibody. The resin-bound proteins were detected with anti-Flag and anti-GFP antibodies. D, BiFC analysis of AtAIRP5/GARU^C208S and AtSCPL1 in N. benthamiana epidermal cells in the presence of MG132. The 35S:Venus^N-Myc-AtAIRP5/GARU^C208S, 35S:AtSCPL1-HA-Venus^C, and 35S:BiP1-mRFP-HDEL (an ER marker) were co-infiltrated into N. benthamiana leaves using the Agrobacterium-mediated method for transient expression. After 2–3 days, the leaves were treated with MG132 (50 µM) for 2 h. Reconstituted Venus signals were visualized by confocal microscopy. The 35S:Venus^N-Myc-AtAIRP5/GARU^C208S + 35S:HA-Venus^C and 35S:Venus^N-Myc + 35S:AtSCPL-HA-Venus^C were used as negative controls. Scale bars = 20 μm.

The interaction between AtAIRP5/GARU and AtSCPL1 was further tested by a bimolecular fluorescence complementation (BiFC) assay. The yellow fluorescent protein Venus was spliced into the N-terminal region (Venus^N) and the C-terminal region (Venus^C). Venus^N and Venus^C were fused to the N- and C-termini of AtAIRP5/GARU^C208S and AtSCPL1, respectively. When Venus^N-Myc-AtAIRP5/GARU^C208S and AtSCPL1-HA-Venus^C were coexpressed in the presence of MG132 (50 µM), an inhibitor of the 26S proteasome complex, in N. benthamiana leaf epidermal cells, web-like network signals were detected throughout the cytosolic fractions (Figure 4D). These reconstituted fluorescent signals were closely overlapped with those of BiP1-mRFP-HDEL, an ER marker protein. However, in the absence of MG132, coexpression of Venus^N-Myc-AtAIRP5/GARU^C208S and AtSCPL1-HA-Venus^C did not give rise to detectable binding signals (Supplemental Figure S8). Interaction of Venus^N-Myc-AtAIRP5/GARU^C208S and AtSCPL1-HA-Venus^C was unchanged in the presence of ABA in addition to MG132 (Supplemental Figure S9). Overall, the results of these in vitro and in vivo interaction analyses indicated that AtSCPL1 interacts with AtAIRP5/GARU.

The AtSCPL1 protein was degraded through the UPS in an AtAIRP5/GARU-dependent manner

The interaction of AtAIRP5/GARU with AtSCPL1 raised the possibility that AtSCPL1 is subject to UPS-dependent degradation in the ABA-mediated drought response. To test this hypothesis, we investigated whether AtSCPL1 turnover is related to the 26S proteasome. The 35S:AtSCPL1-Flag construct was transiently expressed in N. benthamiana leaves. After 2 days of incubation, leaf discs were prepared and then treated with the protein synthesis inhibitor cycloheximide (CHX) for 2–4 h with or without MG132. Then, total proteins were extracted and examined by immunoblotting with an anti-Flag antibody. The results showed that ∼28% (at 2 h) and 64% (at 4 h) of AtSCPL1-Flag was degraded in the absence of MG132 (Figure 5A). However, in the presence of MG132, AtSCPL1-Flag was more stable and ∼84% of the protein was retained, suggesting that the stability of AtSCPL1 was regulated by the 26S proteasome. To determine whether AtSCPL1 is a substrate protein of AtAIRP5/GARU, an in vitro target ubiquitination assay was conducted. Purified 6×His-3×Myc-ΔSP-AtSCPL1 was incubated with MBP-AtAIRP5/GARU or MBP-AtAIRP5/GARU^C208S in the presence or absence of E1 (Arabidopsis UBA1) at 30°C for 1 h, and the reaction products were detected by immunoblotting with anti-Myc and anti-Ub antibodies. As shown in Figure 5B, high molecular mass shifted bands were detected in the lane where 6×His-3×Myc-ΔSP-AtSCPL1 was co-incubated with MBP-AtAIRP5/GARU. In contrast, MBP-AtAIRP5/GARU^C208S was unable to ubiquitinate 6×His-3×Myc-ΔSP-AtSCPL1.

Figure 5.

Regulation of AtSCPL1 protein level by AtAIRP5/GARU in a 26S proteasome-dependent manner. A, CHX chase assay of AtSCPL1. The AtSCPL1-Flag fusion protein was transiently expressed in N. benthamiana leaf epidermal cells. After 2 days of incubation, leaf discs were prepared and treated with the protein synthesis inhibitor CHX (100 μM) for 4 h in the presence or absence of MG132 (50 μM). After 2–4 h, AtSCPL1 protein levels were assessed by immunoblot analysis with an anti-Flag antibody. Rubisco large subunit was detected by Ponceau S staining and was used as a loading control. Error bars are the sd of three biologically independent experiments (n > 5); asterisks indicate statistical significance (**P < 0.01, by one-way ANOVA with post hoc Tukey’s HSD test). B, In vitro target ubiquitination assay. Recombinant 6×His-3×Myc-ΔSP-AtSCPL1 protein was co-incubated with or without UBA1 (E1), MBP-AtAIRP5/GARU, and MBP-AtAIRP5/GARU^C208S. The reaction mixtures were subjected to immunoblotting using anti-MBP and anti-Ub antibodies. Vertical dashed lines indicate the high molecular mass ubiquitinated bands. C, Cell-free degradation assay of ΔSP-AtSCPL1. Purified MBP-ΔSP-AtSCPL1 protein was incubated with crude extracts prepared from N. benthamiana leaves, in which either 35S:GFP-AtAIRP5/GARU or 35S:GFP-AtAIRP5/GARU^C208S was transiently expressed, with or without MG132. MBP-ΔSP-AtSCPL1 levels were examined by immunoblotting with an anti-MBP antibody. D, The time-dependent degradation of MBP-ΔSP-AtSCPL1 presented in C was quantified using the ImageJ software. Error bars are the SD of three biologically independent experiments (n > 5); asterisks indicate statistical significance (**P < 0.01, by one-way ANOVA with post hoc Tukey’s HSD test). E, Cell-free degradation assays of ΔSP-AtSCPL1 testing its dependence on the phosphorylation status of the Tyr-321 residue of AtAIRP5/GARU. Bacterially expressed MBP-ΔSP-AtSCPL1 was incubated with N. benthamiana leaf crude extracts in which 35S:GFP-AtAIRP5/GARU, 35S:GFP-AtAIRP5/GARU^Y321E, 35S:GFP-AtAIRP5/GARU^Y321F, or 35S:GFP-AtAIRP5/GARU^C208S was transiently expressed in the presence or absence of MG132. Numbers below the immunoblots are average values obtained from three independent experiments and reflect the ratio between the control lane and the experimental lane with the control lane normalized to 1.0.

The results of the aforementioned CHX chase and in vitro ubiquitination analyses indicated that AtSCPL1 is a target protein of AtAIRP5/GARU and is controlled via the UPS in an AtAIRP5/GARU-dependent fashion. To test this hypothesis, we conducted cell-free degradation assays. The recombinant MBP-ΔSP-AtSCPL1 protein was incubated with N. benthamiana leaf crude extracts that transiently expressed either 35S:GFP-AtAIRP5/GARU or 35S:GFP-AtAIRP5/GARU^C208S in the presence or absence of MG132. Then, the time-dependent changes in the level of MBP-ΔSP-AtSCPL1 protein were assessed using an anti-MBP antibody. As shown in Figure 5, C and D, the amount of MBP-ΔSP-AtSCPL1 was reduced by ∼33% and 65% after 2.5 and 5 h, respectively, with GFP-AtAIRP5/GARU-expressing crude extracts. Degradation of MBP-ΔSP-AtSCPL1 was inhibited by MG132. In contrast, when MBP-ΔSP-AtSCPL1 was incubated with GFP-AtAIRP5/GARU^C208S crude extract, the protein level remained unchanged regardless of MG132 (Figure 5, C and D).

Nemoto et al. (2017) reported that phosphorylation of the Tyr-321 residue in AtAIRP5/GARU resulted in reduced degradation of the GA receptor GID1. Thus, we investigated whether phosphorylation of AtAIRP5/GARU at Tyr-321 affects AtSCPL1 degradation. When MBP-ΔSP-AtSCPL1 was incubated for 1–2 h with an N. benthamiana leaf crude extract expressing AtAIRP5/GARU^Y321E, a Tyr-321 phosphorylation-mimic form, the protein level was reduced to ∼84% at 1 h and to 72% at 2 h of incubation (Figure 5E). These reduction patterns were similar to those observed with wild-type AtAIRP5/GARU-expressing extract. The abundance of MBP-ΔSP-AtSCPL1, however, decreased more rapidly to ∼66% and 42% after 1 and 2 h incubation, respectively, with AtAIRP5/GARU^Y321F, a phosphorylation-null form. The abundance of MBP-ΔSP-AtSCPL1 was stably maintained in the presence of MG132 in all examined crude extracts (Figure 5E). These results suggest that the degradation of AtSCPL1 is promoted by AtAIRP5/GARU with unphosphorylated Tyr-321.

We next investigated whether the AtSCPL1 protein level is regulated by AtAIRP5/GARU in planta. Using an Arabidopsis leaf protoplast transient expression system, we identified that AtSCPL1-GFP was expressed at 2.2-fold higher level in the atairp5/garu mutant protoplasts than in the wild-type protoplasts (Figure 6, A and B). RT–PCR confirmed that the transcript levels of 35S:AtSCPL1-GFP were consistent in the wild-type and atairp5/garu mutant protoplasts. The level of control protein GFP did not vary between the mutant and wild-type Arabidopsis protoplasts. When 35S:AtSCPL1-GFP was transiently expressed in Arabidopsis protoplasts in the presence of 100 µM ABA, the level of AtSCPL1-GFP was reduced to 75% as compared without ABA (Figure 6, C and D). Based on these in vitro and in vivo results, we concluded that turnover of AtSCPL1 is regulated via the UPS in an AtAIRP5/GARU-dependent manner.

Figure 6.

In planta regulation of AtSCPL1 protein level by AtAIRP5/GARU and ABA. A, Transient expression patterns of AtSCPL1-GFP in the protoplasts prepared from wild-type and atairp5/garu seedlings. The 35S:AtSCPL1-GFP or 35S:GFP construct was transiently expressed in the protoplasts prepared from light-grown wild-type and atairp5/garu seedlings. Protoplasts were examined by immunoblotting with anti-GFP. GFP was used as a negative control. The results of RT-PCR analysis of AtSCPL1-GFP, GFP, and AtACT8 are shown in the lower part to confirm that differences in protein levels are not due to differences in transcript levels. B, Quantification of the AtSCPL1-GFP and GFP protein levels shown in (A) using the ImageJ software. Error bars are the sd of four biologically independent experiments (n > 50); asterisks indicate statistical significance (**P < 0.01, by Student’s t test). C, Transient expression patterns of AtSCPL1-GFP in the protoplasts with or without ABA treatment. The 35S:AtSCPL1-GFP and 35S:GFP constructs were transiently expressed in the protoplasts as described in (A). The protoplasts were incubated with ABA (100 µM) or DMSO for 2 h and were subjected to immunoblotting with anti-GFP. RT–PCR analysis of AtSCPL1-GFP, GFP, and AtACT8 is shown in the lower part. D, Protein levels of AtSCPL1-GFP and GFP shown in (C) were quantified. Error bars are the sd of three biologically independent experiments (n > 50); asterisks indicate statistical significance (*P < 0.05, by Student’s t test).

atscpl1 knockout mutant plants were more sensitive to ABA and more tolerant to drought stress than wild-type Arabidopsis plants

Our results indicated that AtSCPL1 is a substrate of AtAIRP5/GARU; thus, we hypothesized that AtSCPL1 plays a negative role in the ABA response. To test this possibility, T-DNA insertion mutants of AtSCPL1 were obtained, and their phenotypes were analyzed. The atscpl1-1 (SALK_063618) and atscpl1-2 (SALK_087447) mutant lines contained T-DNA insertions in the eighth and first exons, respectively (Supplemental Figure S10A). Homozygous atscpl1-1 and atscpl1-2 progeny were verified using genotyping PCR, and RT–PCR confirmed the absence of full-length AtSCPL1 transcripts in these homozygous plants (Supplemental Figure S10, B and C). Both atscpl1-1 and atscpl1-2 mutant seedlings were more sensitive to ABA than wild-type plants at the germination stage (Figure 7, A and B). Although wild-type and atscpl1 seedlings were phenotypically indistinguishable in the absence of ABA, there were marked phenotypic differences in the presence of 0.9-μM ABA; ∼57% of the wild-type seedlings germinated, while only 33%–37% of the atscpl1 seedlings displayed cotyledon greening. In the presence of 1.2-µM ABA, ∼13% and 4%–6% of the wild-type and mutant plants germinated, respectively (Figure 7, A and B).

Figure 7.

Phenotypic analyses of atscpl1 knockout mutant plants in response to ABA and drought stress. A, ABA-dependent germination assay of atscpl1 mutant plants. Wild-type and atscpl1-1 and atscpl1-2 mutant seeds were germinated with ABA (0, 0.6, 0.9, and 1.2 μM). Scale bars = 0.5 cm. B, Quantitation of the results shown in (A). Error bars are the sd of three biologically independent experiments (n > 25); asterisks indicate statistical significance (*P < 0.05 and **P < 0.01, by one-way ANOVA with post hoc Tukey’s HSD test). C, ABA-induced stomatal movements of wild-type, atscpl1-1, and atscpl1-2 plants. The abaxial part of 4-week-old rosette leaves was incubated with ABA (0, 1.0, and 10.0 μM). Stomatal apertures (width/length) were measured. Scale bars = 10 μm. D, Quantitation of the stomatal apertures in (C). Error bars are the sd of three biologically independent experiments (n > 30); asterisks indicate statistical significance (P < 0.01, by one-way ANOVA with post hoc Tukey’s HSD test). E, Drought tolerant phenotypes of atscpl1-1 and atscpl1-2 mutants as compared to wild-type plants. Light-grown, 18-day-old plants were subjected to drought stress for 13 days. After 3 days of re-watering, the survival of these water-stressed plants was determined. The images were digitally extracted for comparison. Scale bars = 2 cm. F, Measurement of leaf water loss rates. The aerial parts of wild-type, atscpl1-1, and atscpl1-2 plants were detached and the FWs were measured at the indicated time points. Error bars are the SD of three biologically independent experiments (n > 12).

Next, we compared the ABA-mediated stomatal movements of wild-type and atscpl1 mutant plants. The average stomatal apertures of wild-type leaves were reduced from 0.207 ± 0.008 (without ABA) to 0.140 ± 0.008 (with 1.0-µM ABA) and 0.113 ± 0.005 (with 10.0-µM ABA), whereas those of atscpl1 mutant plants were reduced from 0.199 ± 0.004–0.204 ± 0.004 (without ABA) to 0.108 ± 0.006–0.109 ± 0.005 (with 1.0-μM ABA) and 0.092 ± 0.004–0.093 ± 0.006 (with 10.0-μM ABA) (Figure 7, C and D). Thus, the stomatal apertures of atscpl1 mutant leaves closed more rapidly than those of wild-type plants in response to ABA. In addition, the atscpl1-1 and atscpl1-2 mutants displayed increased tolerance to drought stress compared to the wild-type plants. In this experiment, wild-type and atscpl1 plants were grown for 18 days under normal growth conditions. These plants were then exposed to water stress by withholding irrigation for 13 days, and their survival was monitored for 3 days after re-irrigation. As shown in Figure 7E, the survival percentages of wild-type, atscpl1-1, and atscpl1-2 plants were 11.6% (8 out of 69), 63.4% (48 out of 71), and 72.5% (50 out of 69), respectively. In addition, the atscpl1 mutant leaves lost their water content more slowly than the wild-type leaves. After 6 h of incubation at room temperature, the FWs of the wild-type and atscpl1 leaves were decreased to ∼63.6% and 70.0%, respectively, of the starting weights (Figure 7F). The ABA-hypersensitive stomatal closure and drought-tolerant phenotypes of atscpl1 plants were opposite of those of atairp5/garu plants (Figure 1, D and E). These results are consistent with the notion that AtSCPL1 is a negative factor in the ABA-mediated dehydration stress response.

AtSCPL1 is downstream of AtAIRP5/GARU in ABA-dependent seed germination, stomatal closure, and the drought stress response

To examine the epistatic relationship between AtAIRP5/GARU and AtSCPL1, we crossed atairp5/garu-1 and atscpl1-2 single mutant plants and obtained an atairp5/garu-1 atscpl1-2 double knockout mutant line. Genotyping PCR and RT–PCR analyses demonstrated that the homozygous atairp5/garu-1 atscpl1-2 mutant expressed undetectable AtAIRP5/GARU and AtSCPL1 mRNAs (Supplemental Figure S11, A and B). The atairp5/garu-1 atscpl1-2 double mutant progeny exhibited an ABA-hypersensitive germination phenotype, which was similar to that of atscpl1-2, rather than that of atairp5/garu-1. The results also showed that the percentages of germinating wild-type, atairp5/garu-1, atscpl1-2, and atairp5/garu-1 atscpl1-2 seedlings were ∼62%, 85%, 34%, and 48%, respectively, in the presence of 0.9-µM ABA and 15%, 26%, 5%, and 7%, respectively, in the presence of 1.2-µM ABA (Figure 8, A and B). In addition, the atairp5/garu-1 atscpl1-2 double mutant plants displayed ABA-hypersensitive stomatal closure, which was similar to that of atscpl1-2 plants. As shown in Figure 8, C and D, the average stomatal apertures of wild-type, atairp5/garu-1, atscpl1-2, and atairp5/garu-1 atscpl1-2 were 0.141 ± 0.014, 0.176 ± 0.004, 0.113 ± 0.005, and 0.126  ± 0.003, respectively, in the presence of 1.0-μM ABA and 0.118 ± 0.006, 0.157 ± 0.008, 0.096 ± 0.010, and 0.102 ± 0.005, respectively, in the presence of 10.0-μM ABA. These results indicate that AtSCPL1 is downstream of AtAIRP5/GARU in ABA-dependent seed germination and stomatal closure.

Figure 8.

Genetic relationship between AtAIRP5/GARU and AtSCPL1. A, ABA-mediated germination assays. Sterilized seeds of wild-type, atairp5/garu-1 and atscpl1-2 single mutant, and atairp5/garu-1 atscpl1-2 double mutant plants were germinated in the presence of ABA (0, 0.6, 0.9, and 1.2 μM). Scale bars = 0.5 cm. B, Quantitation of the data collected in (A). Error bars are the sd of three biologically independent experiments (n > 25); asterisks indicate statistical significance (*P < 0.05 and **P < 0.01 by one-way ANOVA with post hoc Tukey’s HSD test). C, ABA-mediated stomatal closure of wild-type, atairp5/garu-1 and atscpl1-2 single mutant, and atairp5/garu-1 atscpl1-2 double mutant plants. Scale bars = 10 μm. D, Quantitation of the stomatal apertures in (C). Error bars are the sd of three biologically independent experiments (n > 30); asterisks indicate statistical significance (^**P < 0.01, by one-way ANOVA with post hoc Tukey’s HSD test). E, Drought phenotype assays of wild-type, atairp5/garu-1, and atscpl1-2 single mutant, and atairp5/garu-1 atscpl1-2 double mutant plants. Light-grown, 18-day-old plants were exposed to water deficiency for 13 days. After 3 days of rewatering, the survival of these water-stressed plants was assessed. The images were digitally extracted for comparison. Scale bars = 2 cm. F, Measurement of leaf water content in wild-type, atairp5/garu-1 and atscpl1-2 single mutant, and atairp5/garu-1 atscpl1-2 double mutant plants over time. Error bars are the SD of three biologically independent experiments (n > 12).

Next, we compared the survivability of atairp5/garu-1 and atscpl1-2 single and atairp5/garu-1 atscpl1-2 double mutant plants under water stress. After 13 days of dehydration and subsequent re-watering, the survival percentages of wild-type, atairp5/garu-1, atscpl1-2, and atairp5/garu-1 atscpl1-2 plants were 18.6% (18 out of 97), 0% (0 out of 93), 80.4% (78 out of 97), and 38.5% (37 out of 96), respectively (Figure 8E). Consistently, the detached leaf water loss rate of atairp5/garu-1 atscpl1-2 was more similar to that of atscpl1-2, rather than that of atairp5/garu-1, during the 60–360 min incubation period at room temperature (Figure 8F). Taken together, the phenotypic properties of the atairp5/garu-1 atscpl1-2 double mutant progeny indicate that AtSCPL1 functions downstream of AtAIRP5/GARU RING E3 Ub ligase in ABA-dependent seed germination, stomatal closure, and the drought stress response in Arabidopsis.

Expression levels of AtAIRP5/GARU and AtSCPL1 are inversely correlated with the amounts of endogenous ABA under dehydration stress

We considered the possibility that the opposite phenotypes of atairp5/garu and atscpl1 mutant plants in response to water stress might result from differences in endogenous ABA levels. To investigate this possibility, wild-type, atairp5/garu, and atscpl1 single mutant, and atairp5/garu-1 atscpl1-2 double mutant plants were grown for 3 weeks under normal growth conditions, subjected to dehydration for 9 days, and then mature leaves were detached for ABA measurement. Without drought treatment, the amounts of endogenous ABA in leaf tissues were indistinguishable between wild-type and mutant plants (Figure 9A). However, the accumulation of ABA in response to drought stress was more evident in the atscpl1 plants than in the wild-type and atairp5/garu plants; the amounts of endogenous ABA in the wild-type, atairp5/garu, and atscpl1 leaves were 144.7 ± 17.5 ng g⁻¹ FW, 97.3 ± 8.0–100.8 ± 7.6 ng g⁻¹ FW, and 280.0 ± 39.7–299.1 ± 57.1 ng g⁻¹ FW, respectively (Figure 9A). Thus, under dehydration stress, the expression levels of AtAIRP5/GARU and AtSCPL1 were inversely associated with the amount of ABA. Furthermore, the atairp5/garu-1 atscpl1-2 double mutant leaves contained 216.4 ± 24.9 ng g⁻¹ FW of ABA after water stress, which was more similar to the amount of atscpl1 leaves, rather than the amount of atairp5/garu leaves. Because AtSCPL1 is possibly involved in sinapic acid metabolism in Arabidopsis, we next measured endogenous levels of sinapic acid before and after drought stress. As shown in Figure 9B, before drought stress, endogenous amounts of sinapic acid were very similar in wild-type and mutant leaves. However, after drought treatment, amount of sinapic acid in atairp5/garu leaves (864.7 ± 57.9 nmol g⁻¹ FW) was lower than that in wild-type leaves (1,008.1 ± 27.0 nmol g⁻¹ FW). In contrast, those in atscpl1 single (1,252.3 ± 124.1 nmol g⁻¹ FW) and atairp5/garu-1 atscpl1-2 double (1,235.6 ± 76.2 nmol g⁻¹ FW) mutants were significantly higher compared to wild-type leaves (Figure 9B). Therefore, the opposite phenotypes of atairp5/garu and atscpl1 mutants to drought stress were due to, at least in part, the difference in endogenous ABA and sinapic acid levels. Taken together, our data suggest that AtSCPL1 plays a negative role in ABA-mediated drought stress by altering ABA levels downstream of AtAIRP5/GARU RING E3 Ub ligase.

Figure 9.

Working model of the dual roles of AtAIRP5/GARU E3 Ub ligase in the ABA-mediated drought stress response and GA signaling. A, Changes in endogenous ABA content in rosette leaves in response to drought stress. Wild-type, atairp5/garu, and atscpl1 single mutant, and atairp5/garu-1 atscpl1-2 double mutant plants were grown under well-watered conditions for 3 weeks. Then, half of them were watered, while the others were not watered for a period of 9 days. The amounts of ABA in the leaf tissues were measured using an enzyme immunoassay (Phytodetek). Error bars are the sd of four biologically independent experiments (n > 20); asterisks indicate statistical significance (**P < 0.01, by one-way ANOVA with post hoc Tukey’s HSD test). B, Amounts of sinapic acid in rosette leaves before and after water stress. Sinapic acid was extracted from well-watered and water-stressed leaves of wild-type, atairp5/garu-1, and atscpl1-2 single mutant, and atairp5/garu-1 atscpl1-2 double mutant plants and were quantified by HPLC. Error bars are the sd of four biologically independent experiments (n > 20); asterisks indicate statistical significance (**P < 0.01, by one-way ANOVA with post hoc Tukey’s HSD test). C, Working model of the opposing roles of AtAIRP5/GARU in the ABA and GA signaling pathways. AtAIRP5/GARU functions as a positive regulator of the ABA-mediated drought stress response by promoting the UPS-dependent degradation of AtSCPL1. AtAIRP5/GARU also negatively regulates GA signaling by inducing the turnover of the GA receptor GID1.

Discussion

In this study, loss-of-function mutant lines in AtAIRP5/GARU, encoding a cytosolic- and nuclear-located RING-type E3 Ub ligase, were isolated based on their ABA hyposensitivity during germination (Figure 1). Further phenotypic analyses revealed that the atairp5/garu mutant plants exhibited retarded ABA-dependent stomatal closure and were markedly more susceptible to dehydration stress than wild-type plants (Figure 2). These results indicate that AtAIRP5/GARU has a positive effect on the ABA-mediated drought stress response.

AtAIRP5/GARU was previously reported to function as a negative regulator of GA signaling by inducing degradation of the GA receptor GID1 through the 26S proteasome pathway (Nemoto et al., 2017). Numerous studies have provided evidence that GA and ABA antagonistically mediate diverse subsets of physiological events, including seed dormancy, germination, root elongation, and flowering, through complicated crosstalk networks (Vishal and Kumar, 2018; Shu et al., 2018; Lin et al., 2020). In addition, GA has been shown to participate in the responses to abiotic stresses, such as low temperature, high salinity, and dehydration (Achard et al., 2008; Magome et al., 2008; Colebrook et al., 2014; Plaza-Wüthrich et al., 2016). Most recently, Wang et al. (2020) reported that GAI, a DELLA repressor protein, is involved in the ABA-mediated response to water deficiency by interacting with the ABA-responsive element binding factor ABF2. AtAIRP5/GARU downregulates GA signaling by promoting degradation of GID1 (Nemoto et al., 2017) and upregulates ABA signaling by accelerating turnover of AtSCPL1 (Figures 5 and 6; see below). These results prompted us to speculate that AtAIRP5/GARU is a cellular component of the GA–ABA crosstalk that fine-tunes the cellular and physiological responses of Arabidopsis, which helps plants to endure harsh growth conditions.

A series of in vitro and in vivo interaction assays identified AtSCPL1, a serine carboxypeptidase-like1, as an interacting partner of AtAIRP5/GARU (Figures 3 and 4). ΔSP-AtSCPL1, a signal peptide-deleted mature form of AtSCPL1, was ubiquitinated by AtAIRP5/GARU, and its levels were decreased by the UPS in an AtAIRP5/GARU-dependent manner (Figure 5). An in planta study also showed that the abundance of AtSCPL1 is regulated by AtAIRP5/GARU and ABA (Figure 6). Consistently, atscpl1 single and atairp5/garu-1 atscpl1-2 double loss-of-function mutant progeny were hypersensitive to ABA during both germination and postgermination growth and were markedly more tolerant to drought stress than wild-type plants in an ABA-dependent manner (Figures 7 and 8). Thus, it is very likely that AtSCPL1 is a target protein of AtAIRP5/GARU RING E3 Ub ligase, and hence, it is a negative factor in the ABA-modulated drought stress response.

SCPL family members possess acyltransferase activity and utilize 1-O-β-glucose esters as acyl moiety donors (Milkowski and Strack, 2004; Bontpart et al., 2015). These enzymes are involved in a range of processes, from normal growth and development to defense mechanisms against biotic and abiotic stresses in dicot Arabidopsis and monocot crop rice (Oryza sativa) and oat (Avena strigosa) plants (Lehfeldt et al., 2000; Liu et al., 2008; Mugford et al., 2009; Ciarkowska et al., 2018). Based on their protein sequence homology, Arabidopsis AtSCPL proteins can be grouped into two major clades, I and II, and the first group can be further subdivided into clades IA and IB (Fraser et al., 2005). AtSCPL1 belongs to clade IA (Supplemental Figures S6 and S7). Several clade 1A AtSCPLs, including AtSCPL8, AtSCPL9, AtSCPL10, and AtSCPL19, which utilize sinapoylglucose as an acyl donor, have been identified as sinapoyltransferases in Arabidopsis (Lehfeldt et al., 2000; Shirley et al., 2001; Shirley and Chapple, 2003; Fraser et al., 2007). sng1-1 mutant lines, which are defective in the AtSCPL8 gene, were hyposensitive to ABA during the germination stage (Bi et al., 2017). In contrast, sng2 mutant plants, in which AtSCPL19 was suppressed, displayed ABA-hypersensitive germination. The opposite phenotypes of sng1-1 and sng2 mutants were partially due to the difference in endogenous levels of sinapoylcholine, as the sng1-1 and sng2 mutant plants contained higher and lower sinapoylcholine contents, respectively, than wild-type plants under ABA treatment (Bi et al., 2017). The authors also showed that sinapate-treated plants contained higher levels of glucose-conjugated ABA (ABA-GE) but less free ABA than control plants grown under normal conditions. Furthermore, sinapate treatment resulted in increased transcript levels of UGT71C5, UGT71B6, UGT71B7, and UGT71B8, all of which encode proteins that catalyze the glycosylation of ABA to ABA-GE. These results suggest that sinapic acid and its derivatives alter ABA homeostasis in Arabidopsis. In faba bean (Vicia faba) leaves, sinapic acid inhibits ABA-induced stomatal closure (Purohit et al., 1991). With this background in mind, we postulated that AtSCPL1 might function as an acyltransferase that participates in sinapate ester metabolism, and its reactants or products may affect the homeostasis of ABA. This assumption is supported, at least in part, by the finding that atscpl1 single and atairp5/garu-1 atscpl1-2 double mutant leaves contained higher amounts of endogenous ABA and sinapic acid than wild-type and atairp5/garu mutant leaves in response to drought stress (Figure 9, A and B). In this scenario, ABA- and drought-induced AtAIRP5/GARU E3 Ub ligase promotes the UPS-dependent turnover of AtSCPL1, which, in turn, results in increased endogenous ABA levels and an enhanced response to ABA-mediated drought stress (Figure 9C).

Nemoto et al. (2017) showed that phosphorylation at the Tyr-321 residue of AtAIRP5/GARU inhibited the interaction of AtAIRP5/GARU with GID1 and partially rescued the GID1 protein level. In contrast, nonphosphorylated AtAIRP5/GARU promoted the UPS-dependent proteolysis of GID1, suggesting that the phosphorylation status of AtAIRP5/GARU E3 ligase is critical for its activity. We found that the degradation patterns of AtSCPL1 are also associated, at least in part, with phosphorylation of AtAIRP5/GARU at the Tyr-321 residue. When ΔSP-AtSCPL1 was incubated with a N. benthamiana leaf crude extract of AtAIRP5/GARU^Y321F, a phosphorylation-null form, ΔSP-AtSCPL1 was more rapidly degraded than when incubated with crude extracts of the phosphorylation-mimic AtAIRP5/GARU^Y321E and wild-type (Figure 5E). These results suggested the possibility that dephosphorylation of AtAIRP5/GARU at the Tyr-321 residue suppresses GA signaling by promoting GID1 degradation and, in turn, induces ABA signaling by accelerating AtSCPL1 turnover. This reverse regulation of the GA and ABA signaling pathways by Tyr-321 phosphorylation of AtAIRP5/GARU may enable plants to effectively respond to rapidly fluctuating environmental conditions. Thus, we hypothesized that AtAIRP5/GARU is a key regulator of GA and ABA signaling, and its phosphorylation status on Tyr-321 might function as a molecular switch for the GA-ABA crosstalk network.

Taken together, it is highly plausible that AtSCPL1 functions as a negative factor in the response to ABA-modulated dehydration stress, and, in turn, AtAIRP5/GARU mediates the ubiquitination of AtSCPL1 and positively regulates the water stress response (Figure 9C). However, due to the different subcellular localizations of the proteins, that is, AtAIRP5/GARU in the cytosolic and nuclear fractions and AtSCPL1 at the ER (Figure 3C; Supplemental Figure S3B), how AtAIRP5/GARU interacts with and mediates the ubiquitination of AtSCPL1 should be investigated from a cellular point of view. BiFC assays revealed that AtAIRP5/GARU and AtSCPL1 interact each other at the ER in the presence of MG132 with and without ABA (Figure 4D; Supplemental Figure S9). Because the ubiquitination of AtSCPL1 by AtAIRP5/GARU resulted in its degradation via the 26S proteasome (Figures 5 and 6), which is predominantly localized and functions in the cytosol and nucleus (Collins and Goldberg, 2017), we speculated that AtAIRP5/GARU binds to and ubiquitinates AtSCPL1 in the ER, which faces the cytosolic space. Because interaction between AtAIRP5/GARU and AtSCPL1 was detected regardless of ABA, it seems likely that interaction and ubiquitination of AtSCPL1 is mainly dependent on the cellular abundance of AtAIRP5/GARU protein. This assumption is consistent with the notion that the effect of ABA was not as evident as that of AtAIRP5/GARU on the decrease in AtSCPL1 level in the protoplast system (Figure 6). Alternatively, we considered the possibility that the interaction between AtAIRP5/GARU and AtSCPL1 in planta requires an as-yet-unidentified cellular factor(s), which needs to be clarified.

Because AtAIRP5/GARU is a negative regulator of GA signaling (Nemoto et al., 2017), it is possible that its target protein AtSCPL1 is a positive factor in GA signaling. However, atscpl1 mutant lines displayed similar phenotypes compared with wild-type plants in hypocotyl length in the presence or absence of exogenously applied 1-µM paclobutrazol (PAC), a GA inhibitor, and 1- to 5-µM GA₃ (Supplemental Figure S12), suggesting that AtSCPL1 might not be involved in the response to GA.

In conclusion, our results suggest that AtAIRP5/GARU RING E3 Ub ligase plays a negative role in GA signaling and a positive role in ABA signaling in Arabidopsis (Figure 9C). The dual role of AtAIRP5/GARU in the degradation of GID1 and AtSCPL1 could be interconnected as a molecular switch for the cellular balance between GA and ABA signaling, which helps Arabidopsis effectively deal with water-stressed growth conditions.

Materials and methods

Plant materials and growth conditions

Arabidopsis (A.thaliana) ecotype Columbia-0 (Col-0) was used as the wild-type and T-DNA insertion mutant background. The atairp5/garu-1 (SALK_037121), atairp5-2/garu-2 (SALK_127503), atairp5/garu-3 (SALK_137684), atscpl1-1 (SALK_063618), and atscpl1-2 (SALK_087447) mutant seeds were obtained from the Arabidopsis Biological Resource Center (http://abrc.osu.edu). The T-DNA insertion and null mutation of AtAIRP5/GARU were confirmed by genotyping PCR and RT–PCR, respectively. The atairp5/garu-1 atscpl1-2 double knockout mutant plants were generated by genetic crossing of atairp5/garu-1 and atscpl1-2 single mutant plants. The 2×Flag-AtAIRP5/GARU fusion gene was cloned into the pEG100 vector, and the resulting construct was transformed into a wild-type plant via the floral dipping method (Clough and Bent, 1998). The transformed plants were selected using glufosinate-ammonium (BASTA, 30 μg mL⁻¹; Duchefa Biochemie, Haarlem, the Netherlands), and the homozygous T3 transgenic line was used for phenotypic analysis. Ectopic expression of AtAIRP5/GARU transcript and protein in the 2×Flag-AtAIRP5/GARU-overexpressing plants was verified by RT–PCR and immunoblotting with an anti-Flag antibody (Sigma-Aldrich, St Louis, MO, USA), respectively, as described by Kim et al. (2019). Wild-type and transgenic seeds were grown on Murashige and Skoog (MS) medium (Duchefa Biochemie) containing 0%–1% (w/v) sucrose and 0.7% (w/v) phytoagar or on soil (Sunshine mix 5; Sun Gro Horticulture, Agawam, MA, USA) at 22°C under continuous light.

RT–PCR and RT–qPCR analyses

Light-grown, 8-day-old wild-type seedlings were exposed to dehydration stress by uncovering the lid of a plant culture dish for 4 h in a growth chamber (22°C). For the ABA and NaCl treatments, wild-type seedlings were transferred to 0.5× MS liquid medium supplemented with 100-µM ABA (Sigma-Aldrich) or 300-mM NaCl and then incubated for 2 h. Total RNA was extracted from abiotic stress- and ABA-treated seedlings using the Easy Spin Plant RNA Extraction Kit (Intron Biotechnology, Seoul, South Korea). cDNA was synthesized using the SuperScript IV First-Strand cDNA Synthesis Kit (ThermoFisher, Waltham, MA, USA) according to the manufacturer’s protocol. RT–qPCR was conducted as described by Ryu et al. (2010). Transcript levels were normalized to that of the glyceraldehyde-3-phosphate dehydrogenase C subunit as an internal control. The primers used in this study are listed in Supplemental Table S1.

Germination assay, stomatal aperture measurement, and phenotypic analyses in response to ABA and drought stress

For the ABA-mediated germination assay, wild-type and mutant seeds were sterilized and incubated at 4°C for imbibition. After 48 h, the seeds were sown on MS solid medium (Duchefa Biochemie) containing various concentrations (0.6, 0.9, and 1.2 µM) of ABA (Sigma Aldrich) and incubated at 22°C under continuous light conditions in a growth chamber. The percentages of cotyledon greening were calculated at 10 days after sowing as described by Kim and Kim (2013).

Stomatal aperture measurements were conducted as described previously (Cho et al., 2011; Seo et al., 2012), with a slight modification. In brief, the abaxial part of the mature rosette leaves of 4-week-old plants was peeled off and incubated in stomatal opening solution (10-mM KCl, 100-µM CaCl₂, and 10-mM MES, pH 6.1; Kwak et al., 2003). After incubation for 4 h, the leaves were transferred to the same solution containing different concentrations (0, 1.0, and 10.0 µM) of ABA for 2 h. At least 30 stomatal apertures (width/length) on each leaf were measured using the ImageJ software (http://imagej.nih.gov/ij/).

For the drought phenotype analysis, wild-type, atairp5/garu, and atscpl1 single, and atairp5/garu atscpl1 double mutant plants were grown for 18 days in soil under continuous light conditions and then exposed to water deficiency for 11–13 days. After 3 days of re-watering, the survival percentages of the plants were determined. A leaf water loss assay was conducted as described by Yu et al. (2020).

In vitro pull-down and in vivo co-IP assays

An in vitro pull-down assay was carried out as described by Kim et al. (2017), with slight modifications. Bacterially expressed 6×His-3×Myc-ΔSP-AtSCPL1 recombinant protein (500 ng) was co-incubated with MBP-AtAIRP5/GARU or MBP (500 ng) in the presence of Ni-NTA agarose resin (Qiagen, Hilden, Germany) in pull-down buffer [1× PBS and 0.5% (v/v) Triton X-100] for 2 h at 4°C. After washing 3 times with pull-down buffer, the resin-bound proteins were eluted with 4× SDS sample buffer.

An in vivo co-IP experiment was conducted according to the method described by Seo et al. (2016). The 35S:AtSCPL1-GFP and 35S:GFP constructs were coexpressed in the presence or absence of 35S:2×Flag-AtAIRP5/GARU + 35S:p19 in N. benthamiana leaves using the Agrobacterium-mediated infiltration method. After 48 h, the infiltrated leaves were ground in liquid nitrogen, and total proteins were extracted in extraction buffer (50-mM Tris–HCl, pH 7.4, 10-mM MgCl₂, 1-mM EDTA, and 0.05% Triton X-100). Leaf extracts (containing 500 µg of total proteins) were incubated with anti-Flag M2 magnetic beads (Sigma Aldrich) for 2 h at 4°C and washed 3 times with extraction buffer. The precipitated proteins were detected by immunoblot analysis using anti-Flag (Sigma Aldrich) and anti-GFP (Clontech, Mountain View, CA, USA) antibodies.

Yeast two-hybrid assays

Yeast two-hybrid analysis was performed according to the protocol described by Kim and Kim (2013). Briefly, the full-length coding region of AtAIRP5/GARU was cloned into the pGBKT7 vector. Then, the generated AtAIRP5/GARU-pGBKT7 plasmid (as bait) and an Arabidopsis cDNA library of 3-day-old etiolated seedlings (as prey) were co-transformed into Saccharomyces cerevisiae strain AH109 (Clontech) and cultured in SD/Trp/–Leu/–His medium for 3 days at 30°C. To examine the interaction between AtAIRP5/GARU and AtSCPL1, the AtAIRP5/GARU-pGBKT7 construct was co-transformed along with AtSCPL1-pGADT7 or ΔSP-AtSCPL1-pGADT7 into AH109 cells and incubated for 3 days. p53 + T-antigen and lambda + T-antigen were used as positive and negative controls, respectively.

CHX chase, in vitro ubiquitination, and cell-free degradation assays

For the CHX chase experiment, the 35S:AtSCPL1-Flag fusion gene was transiently expressed in N. benthamiana leaves. After 2 days, the leaf discs were incubated in 0.5× MS liquid medium with 100-μM CHX (Sigma Aldrich) in the presence or absence of 50-μM MG132 (AG Scientific) for 2–4 h. Then, AtSCPL1-Flag protein levels were examined by immunoblot analysis with an anti-Flag antibody and quantified using the ImageJ software (https://imagej.nih.gov/).

For the in vitro self-ubiquitination assay, bacterially expressed MBP-AtAIRP5/GARU and MBP-AtAIRP5/GARU^C208S recombinant proteins (300 ng) were incubated with Ub, ATP, E1 (AtUBA1), and E2 (AtUBC8) at 30°C for 1.5 h in ubiquitination buffer (50-mM Tris–HCl, pH 7.5, 2.5-mM MgCl₂, 4-mM ATP, and 0.5-mM DTT) as described by Yu et al. (2021). Purified 6×His-3×Myc-ΔSP-AtSCPL1 recombinant protein (300 ng) was subjected to an in vitro target ubiquitination assay according to the method described by Kim et al. (2017). The ubiquitinated proteins were detected by immunoblotting with anti-MBP (Applied Biological Materials, Richmond, Canada), anti-Myc (Abcam, Cambridge, UK), and anti-Ub (Agrisera, Vännäs, Sweden) antibodies.

Cell-free crude extracts were prepared from N. benthamiana leaves expressing the 35S:GFP-AtAIRP5/GARU + 35S:p19, 35S:GFP-AtAIRP5/GARU^C208S + 35S:p19, 35S:GFP-AtAIRP5/GARU^Y321F + 35S:p19, and 35S:GFP-AtAIRP5/GARU^Y321E + 35S:p19 fusion genes using extraction buffer [1× PBS, 10-mM MgCl₂, and 0.1% (v/v) Triton X-100], as described previously (Seo et al., 2016). The MBP-ΔSP-AtSCPL1 recombinant protein (300 ng) was incubated with the crude extracts (containing 50 µg of total proteins) with or without 50-μM MG132 for 1–5 h. Rubisco large subunit was used as a loading control. Protein bands were quantified using the ImageJ software (https://imagej.nih.gov/).

In planta protein degradation assays in Arabidopsis protoplasts

For AtSCPL1-GFP and GFP stability assays, the protoplasts prepared from light-grown 7-day-old wild-type and atairp5/garu seedlings were transfected with pBI221 harboring the 35S: AtSCPL1-GFP or 35S:GFP construct using the PEG-mediated transfection method (Min et al., 2019). After a 16-h incubation at 22°C under the condition of continuous light, the protoplasts were subjected to immunoblot analysis with anti-GFP antibody (Clontech).

For ABA treatment, protoplasts of wild-type seedlings were transfected with pBI221 harboring the 35S:AtSCPL1-GFP or 35S:GFP construct. After 13 h, the transfected protoplasts were further incubated with 100-μM ABA or dimethyl sulfoxide (DMSO) for 2 h and analyzed by immunoblotting with anti-GFP antibody. Protein bands were quantified using the ImageJ software (https://imagej.nih.gov/).

Subcellular localization

Agrobacterium tumefaciens strain GV3101 cells, which contained 35S:p19 + 35S:AtAIRP5/GARU^C208S-GFP + 35S:mRFP or 35S:19 + 35S:AtSCPL1-GFP + 35S:BiP1-mRFP-HDEL, were infiltrated into 2-week-old N. benthamiana leaves, as described by Son et al. (2010). After 2–3 days of incubation, GFP and red fluorescent protein (RFP) fluorescent signals were detected by confocal microscopy (LSM-880; Carl Zeiss, Oberkochen, Germany). The laser signals were measured using a sliding 76–78 nm detection window. The GFP and RFP signals were acquired with 488- and 561-nm excitation wavelengths, respectively.

BiFC assay

The full-length coding region of AtAIRP5/GARU^C208S was cloned in the pDEST-VYNE(R) vector that contained the N-terminal region (Venus^N; 1 to −173 amino acids) of Venus, as described by Gehl et al. (2009). The full-length coding region of AtSCPL1 was introduced into the pDEST-VYCE vector containing the C-terminal region (Venus^C: 156–239 amino acids) of Venus. The Venus^N-Myc-AtAIRP5/GARU^C208S and AtSCPL1-HA-Venus^C constructs were co-infiltrated into the N. benthamiana leaves. After 2–3 days, the leaf discs were incubated with or without 50-μM MG132 (Sigma Aldrich) containing 1/2× MS medium in the presence or absence of 50-μM ABA (Sigma Aldrich) for 2 h. The reconstituted fluorescent signals were visualized by confocal microscopy (LSM-800). The laser signals were measured using a sliding 69- to 44-nm detection window. The Venus and RFP signals were acquired with 495- and 590-nm excitation wavelengths, respectively. Expression of Venus^N-Myc-AtAIRP5/GARU^C208S and AtSCPL1-HA-Venus^C protein was examined by immunoblotting with anti-Myc (Abcam) and anti-HA (Applied Biological Materials) antibodies. The 35S:Venus^N-Myc-AtAIRP5/GARU^C208S + 35S:HA-Venus^C and 35S:Venus^N-Myc + 35S:AtSCPL1-HA-Venus^C were used as negative controls.

Measurement of ABA and sinapic acid content

ABA content was measured as described by Yang et al. (2015), with slight modifications using the Phytodetek ABA enzyme immunoassay test kit (Agdia, Elkhart, IN, USA). Wild-type, atairp5/garu-1, atairp5/garu-3, atscpl1-1, and atscpl1-2 single, and atairp5/garu-1 atscpl1-2 double mutant plants were grown in soil for 3 weeks and exposed to drought stress for 9 days, and then rosette leaves were collected to measure ABA content. Frozen samples were ground in liquid nitrogen and extracted in 80% (v/v) methanol with 10-mg L⁻¹ butylated hydroxytoluene and 50-mg L⁻¹ citric acid overnight in the dark at 4°C. After centrifugation at 1,000g for 20 min at 4°C, the supernatant was collected and diluted 10-fold with 1× Tris-buffered saline (TBS) buffer (50-mM Tris buffer, pH 7.5, 150-mM NaCl, and 1-mM MgCl₂). The diluted samples were evaluated using the ABA enzyme immunoassay test kit, according to the manufacturer’s protocol.

To measure sinapic acid content in mock- and drought-treated rosette leaves, sinapic acid was extracted with 80% (v/v) methanol at 4°C. After overnight incubation, the samples were centrifuged at 13,000g for 10 min, and the supernatants were subjected to high performance liquid chromatography (HPLC) (Agilent Technologies, Santa Clara, CA, USA). The sample was injected into a Kinetex 2.6 μm C18 Column (50 × 2.1 mm) at a flow of 0.2 mL min⁻¹ using 45% acetonitrile containing 0.2% acetic acid for 4 min. During 4–8 min, acetonitrile concentration increased from 45% to 100%. UV absorption was monitored at 330 nm. The peaks of sinapic acid were checked and quantified using commercially available sinapic acid and chrysin as an internal standard.

Hypocotyl elongation assay with or without PAC and GA treatments

Stratified seeds of wild-type and atscpl1 mutant plants were germinated on Gamborg’s B5 medium in the growth chamber at 23°C under a 16-h light/8-h dark cycle. After 5 days, the seedlings were transferred on 1/2× MS medium supplemented with or without 1-µM PAC, a GA inhibitor, and 1–5 µM GA₃ (Sigma Aldrich) and further incubated for 5 days under dark condition. Seedlings were photographed and hypocotyl length was measured using the ImageJ software (https://imagej.nih.gov/).

Sequence alignment and phylogenetic analyses

The amino acid sequences were aligned using ClustalW software in the BioEdit program (https://bioedit.software.informer.com/). The multiple alignment results were generated, and phylogenetic trees were constructed using the Neighbor-Joining method in MEGA-X software. In phylogenetic trees, branch lengths indicated the evolutionary distance, which was feasible to deduce with the scale of each phylogenetic tree.

Accession numbers

Sequence data from this article can be found in the GenBank/EMBL data libraries under accession numbers AtAIRP5/GARU (At2g40830), AtSCPL1 (At5g36180), AtSCPL2 (At1g73300), AtSCPL6 (At1g73270) AtSCPL8 (At2g22990), AtSCPL10 (At2g23000), AtSCPL9 (At2g23010), and AtSCPL19 (At5g09640).

Supplemental data

The following materials are available in the online version of this article.

Supplemental Figure S1. Identification of homozygous T-DNA insertion mutant lines of AtAIRP5/GARU.

Supplemental Figure S2. Sequence analysis of AtAIRP5/GARU.

Supplemental Figure S3. AtAIRP5/GARU is a cytosolic- and nuclear-localized RING E3 Ub ligase.

Supplemental Figure S4. Phenotypic analyses of AtAIRP5/GARU-overexpressing transgenic Arabidopsis plants in response to ABA during the germination stage.

Supplemental Figure S5. Phenotypic analyses of 35S:2×Flag-AtAIRP5/GARU-atairp5/garu complementation lines in response to ABA during germination.

Supplemental Figure S6. Phylogenetic analysis of the 19 AtSCPL proteins belonging to clade 1A.

Supplemental Figure S7. Amino acid sequence alignment of AtSCPL family members in Arabidopsis.

Supplemental Figure S8. BiFC analysis of Venus^N-Myc-AtAIRP5/GARU^C208S and AtSCPL1-HA-Venus^C without MG132 in N. benthamiana epidermal cells.

Supplemental Figure S9. BiFC analysis of Venus^N-Myc-AtAIRP5/GARU^C208S and AtSCPL1-HA-Venus^C in the presence of ABA in addition to MG132 in N. benthamiana epidermal cells.

Supplemental Figure S10. Identification and molecular characterization of T-DNA-inserted loss-of-function mutant alleles of AtSCPL1.

Supplemental Figure S11. Construction and molecular characterization of atairp5/garu-1 atscpl1-2 double mutant line.

Supplemental Figure S12. Effects of exogenously applied PAC, a GA inhibitor, and GA₃ on the hypocotyl length of wild-type and atscpl1 mutant seedlings.

Supplemental Table S1. Sequences of PCR primers used in this study.

 

N.H.C., O.G.W., E.Y.K., K.Y.P., D.H.S., S.G.Y., Y.A.C., and  J.-H.L. performed the experiments. N.H.C., O.G.W., D.H.S.,  J.-H.L., and W.T.K. analyzed the data. N.H.C., O.G.W., J.-H.L., and W.T.K. planned the project, and N.H.C.,  J.-H.L., and W.T.K. wrote the article. W.T.K. supervised the project and complemented the writing.

The author responsible for distribution of materials integral to the findings presented in this article in accordance with the policy described in the Instructions for Authors (https://academic.oup.com/plphys/pages/general-instructions) is Woo Taek Kim (wtkim@yonsei.ac.kr).

Funding

This work was supported by grants from the National Research Foundation (Mid-Career Researcher Program Project No. 2017R1A2B2006750 and Basic Science Research Program Project Number 2018R1A6A1A03025607), Republic of Korea, to W.T.K.

Conflict of interest statement. The authors declare no conflict of interest.