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. 2020 Dec 16;229(4):2035–2049. doi: 10.1111/nph.17001

COP1 promotes ABA‐induced stomatal closure by modulating the abundance of ABI/HAB and AHG3 phosphatases

Qingbin Chen 1,*, Ling Bai 1,*, Wenjing Wang 1,*, Huazhong Shi 2, José Ramón Botella 3, Qidi Zhan 1, Kang Liu 1, Hong‐Quan Yang 4, Chun‐Peng Song 1,✉
PMCID: PMC7898331  PMID: 33048351

Summary

  • Plant stomata play a crucial role in leaf function, controlling water transpiration in response to environmental stresses and modulating the gas exchange necessary for photosynthesis. The phytohormone abscisic acid (ABA) promotes stomatal closure and inhibits light‐induced stomatal opening. The Arabidopsis thaliana E3 ubiquitin ligase COP1 functions in ABA‐mediated stomatal closure. However, the underlying molecular mechanisms are still not fully understood.

  • Yeast two‐hybrid assays were used to identify ABA signaling components that interact with COP1, and biochemical, molecular and genetic studies were carried out to elucidate the regulatory role of COP1 in ABA signaling.

  • The cop1 mutants are hyposensitive to ABA‐triggered stomatal closure under light and dark conditions. COP1 interacts with and ubiquitinates the Arabidopsis clade A type 2C phosphatases (PP2Cs) ABI/HAB group and AHG3, thus triggering their degradation. Abscisic acid enhances the COP1‐mediated degradation of these PP2Cs. Mutations in ABI1 and AHG3 partly rescue the cop1 stomatal phenotype and the phosphorylation level of OST1, a crucial SnRK2‐type kinase in ABA signaling.

  • Our data indicate that COP1 is part of a novel signaling pathway promoting ABA‐mediated stomatal closure by regulating the stability of a subset of the Clade A PP2Cs. These findings provide novel insights into the interplay between ABA and the light signaling component in the modulation of stomatal movement.

Keywords: abscisic acid (ABA), clade A PP2C, COP1, stomatal movement, ubiquitination

Introduction

Stomata, consisting of guard cells, are highly specialized epidermal structures that regulate gas exchange and water transpiration in terrestrial plants. Stomatal aperture is modulated through changes in the intracellular turgor of guard cells to balance the uptake of CO2 for photosynthesis and maintain water status in response to environmental conditions such as light intensity, water availability, CO2 concentration, etc. (Assmann, 1993; Blatt, 2000; Schroeder et al., 2001; Shimazaki et al., 2007). The stomatal response to these environmental conditions is often mediated by the phytohormone abscisic acid (ABA), and the underlying molecular mechanisms have been extensively studied (Kim et al., 2010).

Abscisic acid promotes stomatal closure and inhibits light‐induced stomatal opening. Perception of ABA in guard cells is dependent on the pyrabactin resistance (PYR)/PYR1‐like (PYL)/regulatory components of ABA receptor (RCAR) protein family of ABA receptors (Lee et al., 2013). The binding of ABA to PYR/PYL/RCAR receptors recruits type 2C protein phosphatase (PP2C) co‐receptors, resulting in the release of PP2Cs from the inhibitory PP2Cs‐Open stomata 1 (OST1) complex (Ma et al., 2009; Xie et al., 2012; Wang et al., 2018). The kinase activity of OST1, an SNF1‐related protein kinase 2 (SnRK2), is inhibited when complexed with PP2Cs, but the recruitment of clade A PP2Cs upon ABA binding with the ABA receptor de‐represses OST1 activity (Soon et al., 2012). The activated OST1 then phosphorylates downstream targets, including the plasma membrane NADPH oxidase and Ca2+ channels, inducing the subsequent accumulation of second messengers such as reactive oxygen species (ROS) and Ca2+ in the guard cells (Song et al., 2014). These second messengers trigger signaling pathways leading to the physiological responses to ABA (Kim et al., 2010; Kollist et al., 2014; Zhang et al., 2014). In addition, OST1‐mediated phosphorylation activates the S‐type slow anion channel‐associated 1 (SLAC1) and the R‐type quickly‐activating anion channel 1 (QUAC1) but inactivates the K+ inward rectifying channel (KAT1) in Arabidopsis. These phosphorylation events cause a massive outflow of NO3 − and Cl− anions from the guard cells, while inhibiting K+ influx into them. As a result, the turgor pressure of the guard cells decreases and the stomata close (Lee et al., 2009; Sato et al., 2009; Imes et al., 2013).

In contrast to ABA, light is one of the most important environmental signals that induces stomatal opening. Stomatal opening during the day is mainly dependent on the coordination of red and blue light (Shimazaki et al., 2007). High‐intensity red light induces stomatal opening by stimulating the production of osmotic metabolites, thus maintaining the turgor of guard cells (Ogawa et al., 1978; Shimazaki et al., 2007). However, the pathway used by red light receptors in guard cells to regulate stomatal movement remains elusive. Compared to red light, which requires a prolonged time period for the induction of stomatal opening, blue light induces stomatal opening in a much faster way. Upon perception by the receptors, blue light quickly induces the phosphorylation of type 1 protein phosphatase (PP1), type 7 protein phosphatase (PP7) and blue light signaling 1 (BLUS1), promoting H+‐ATPase activity on the cell membrane of guard cells (Takemiya et al., 2006; Sun et al., 2012; Takemiya et al., 2013). The activated H+‐ATPase results in a massive outflow of H+, leading to hyperpolarization of the guard cell membrane. The elevated membrane potential activates the voltage‐gated K+ channels, and the resulting K+ influx increases the turgor of the guard cells, which promotes stomatal opening (Shimazaki et al., 2007). Although light and ABA regulate stomatal movement by modulating the turgor pressure of guard cells, the mechanistic relationship between light and ABA signaling in stomatal movement is not fully understood.

CONSTITUTIVELY PHOTOMORPHOGENIC 1 (COP1) is a master negative regulator of photomorphogenesis and plays an important role in the transduction of red, far‐red, blue and ultraviolet (UV) light signals (Lau & Deng, 2012). COP1 is an E3 ubiquitin ligase composed of a RING domain, a coiled helix domain and a WD40 domain, and mediates the ubiquitination and subsequent degradation of several regulatory factors in plant photomorphogenesis (Torii et al., 1998). In dark conditions, COP1 and Suppressor of phya‐105 (SPA) form hetero‐complexes in the nucleus, mediating the ubiquitination of transcription factors such as Elongated hypocotyl 5 (HY5), HY5 homologue (HYH), Long hypocotyl in far‐red light 1 (HFR1), and Long after far‐red light 1 (LAF1) for degradation, thus inhibiting photomorphogenesis and promoting skotomorphogenesis in plants (Osterlund et al., 2000; Holm et al., 2002; Seo et al., 2003; Duek et al., 2004; Balcerowicz et al., 2017). In the light, COP1 is exported from the nucleus to the cytosol, resulting in the accumulation of these transcription factors and promoting photomorphogenesis (Podolec & Ulm, 2018). In recent decades, the functions of COP1 in light signaling have been extensively studied, and accumulating evidence has shown that COP1 is also involved in flowering, biological clock rhythm, virus defense, plant hormone signaling, stomatal development and closing, and other biological processes (Mao et al., 2005; Kang et al., 2009; Wang et al., 2019). However, the role of COP1 in ABA signaling remains unexplored.

In this study, we demonstrate that COP1 is involved in ABA‐mediated stomatal closure. In cop1 mutants, impaired ABA‐promoted stomatal closure is evident, and genetic analyses indicate that COP1 controls stomatal movement upstream of the clade A PP2Cs, which function as ABA signaling regulators. We show that COP1 ubiquitinates ABI/HAB and AHG3 PP2Cs, triggering their degradation, and the COP1 ubiquitination of these PP2Cs is enhanced by ABA. The COP1‐mediated ubiquitination and degradation of the PP2Cs, which are key players in ABA signaling, provides a novel pathway in the modulation of stomatal closing.

Materials and Methods

Plant materials and growth conditions

All the genotypes used in this study were generated in the Columbia (Col‐0) ecotype background. The mutants used in this study include cop1‐4 (Ma et al., 2002), cop1‐6 (Ma et al., 2002), and GUS‐COP1 (Osterlund & Deng, 1998). The T‐DNA insertion mutants abi1‐3 (SALK_076309) and ahg3‐1 (SALK_028132) were obtained from the Arabidopsis Biological Resource Center (ABRC; http://abrc.osu.edu/). cop1‐4abi1‐3, cop1‐4ahg3‐1, and cop1‐4abi1‐3ahg3‐1 mutants were generated by genetic crossing. The primers for identification of these mutants are listed in Supporting Information Table S1. The cop1‐4 plants expressing 35S:ABI1‐MYC or 35S:AHG3‐MYC were generated by crossing the 35S:ABI1‐MYC or 35S:AHG3‐MYC transgenic plants with cop1‐4 mutants. The seedlings were grown in a 1 : 1 mixture of forest soil and vermiculite under 150 μmol photons m−2 s−1 illumination at 22°C with a 12 h : 12 h, light : dark photoperiod in a glasshouse.

Measurements of stomatal aperture

Measurements of stomatal aperture were performed as described previously (Wang et al., 2020). Briefly, epidermal peels were stripped from leaves of 4‐wk‐old plants grown in a glasshouse under the conditions described in the previous paragraph, and incubated in the resting buffer for 2.5 h under a light intensity of 150 μmol m−2 s−1 or in the dark with or without 1 μM ABA. This experiment was repeated three times, and four to six independent plants were used in each experiment.

Bimolecular fluorescence complementation (BiFC) analysis

The coding sequences of the PP2Cs (ABI1, ABI2, HAB1, HAB2, AHG3) with the N‐terminal fragments of GFP (nGFP‐COP1) and COP1 with the C‐terminal fragments of GFP (cGFP‐COP1) were cloned into the pCAMBIA1300 vector to create fusion genes. The PP2C family member PP2C5, which is incapable of interacting with COP1, was used as a negative control, and the COP1‐interacting protein HY5 was used as a positive control (Kudla & Bock, 2016). The combinations of cGFP‐COP1 with nGFP‐PP2Cs were co‐transformed into Nicotiana benthamiana leaves. H2B‐mCherry was expressed in N. benthamiana leaves as a marker protein for nuclear localization (Guo et al., 2018). Three days after the first injection, a second injection was carried out with infiltration buffer containing 50 μM MG132. The tobacco leaves were then cultured for 12 h, and the fluorescence images were captured using a Zeiss LSM 710 scan confocal microscope (Heidenheim, Germany). Primers used for the BiFC assays are listed in Table S1.

Co‐immunoprecipitation (Co‐IP) assay in Arabidopsis protoplasts

The coding sequences of COP1 with a CFP tag (CFP‐COP1) and PP2Cs (ABI1, ABI2, HAB1, HAB2, AHG3) with an MYC tag (PP2Cs‐MYC) were cloned into the pCAMBIA1300 vector to create fusion genes, and these plasmids were co‐transformed into Arabidopsis protoplasts. The protoplasts were lysed by sonication in protein extraction buffer containing 50 mM Tris‐HCl (pH 7.4), 150 mM NaCl, 10 mM MgCl2, 0.2% (v/v) glycerol, 0.1% (v/v) Nonidet P‐40, 1 mM phenylmethylsulfonyl fluoride (PMSF) and 1 × complete protease inhibitor (Roche), and then centrifuged at 12 000 g for 15 min at 4°C. The supernatant was incubated with 15 μl anti‐GFP antibody‐conjugated agarose (Chromotek) for 3 h at 4°C. The Co‐IP products were washed briefly with extraction buffer three times at 4°C, and immune blots with anti‐MYC antibody and anti‐GFP antibody were carried out. To test the effect of ABA on the interactions between COP1 and PP2Cs, a final concentration of 0, 1, or 10 μM ABA was added to the supernatant incubated with anti‐GFP antibody‐conjugated agarose. Primers used for Co‐IP assays are listed in Table S1.

Semi‐in vivo protein degradation analysis

Semi‐in vivo protein degradation analysis was performed as described previously (Liu et al., 2010). Tobacco leaves were injected with Agrobacterium carrying CFP‐COP1, ABI1‐MYC or AHG3‐MYC. The leaves without the Agrobacterium injection (wild‐type, WT) were also collected as controls. Proteins were extracted from the leaves using the extraction buffer described in the subsection ‘Co‐immunoprecipitation (Co‐IP) assay in Arabidopsis protoplasts’, above. A final concentration of 10 μM ATP was added to the cell lysates to maintain the function of the 26S proteasome. The extract containing ABI1‐MYC was mixed with the extract containing CFP‐COP1 or WT extract at a ratio of 1 : 1, and the extract containing AHG3‐MYC was also mixed with the extract containing CFP‐COP1 or WT extract at 1 : 1 ratio. These mixtures were either supplemented with 50 μM MG132 or not supplemented. The mixture was incubated at room temperature and sampled at 0, 2, 4 and 6 h.

For semi‐in vivo degradation analysis of ABI1‐MYC and AHG3‐MYC in WT and pyr1pyl1pyl2pyl4pyl5pyl8 (hexpyl) backgrounds, 10 μg Pro35S:CFP‐COP1 plasmid and 10 μg Pro35S:ABI1‐MYC plasmid, 10 μg Pro35S:CFP‐COP1 plasmid and 10 μg Pro35S:AHG3‐MYC plasmid were mixed and co‐transformed into WT and hexpyl mutant protoplasts respectively. After incubation for 16 h, total proteins were extracted using the extraction buffer and divided into three groups: one without ABA treatment, one with 10 μM ABA and the other with 10 μM ABA plus 50 μM MG132. The total proteins were incubated at room temperature and sampled at 0, 4.5 and 9 h.

In vitro ubiquitination assays

In vitro ubiquitination assays were performed as described previously (Saijo et al., 2003; Liu et al., 2010; Xu et al., 2014), and GST‐ABI1 and GST‐AHG3 proteins were acquired using these methods. The primers for GST‐ABI1 and GST‐AHG3 are listed in Table S1. CFP and CFP‐COP1 samples were extracted from N. benthamiana leaves in which they were transiently expressed. CFP was used as a negative control. The cell lysates were then mixed and immunoprecipitated with 15 μl anti‐GFP antibody‐conjugated agarose. The immunoprecipitated products were washed briefly with extraction buffer three times at 4°C. The ubiquitination reaction mixtures (60 μl) contained 30 ng of UBE1 (E1; Boston Biochem, Cambridge, MA, USA), 30 ng of UbcH5b (E2; Boston Biochem), 10 μg of HA‐tagged ubiquitin (HA‐Ub; Boston Biochem), 200 ng of GST‐ABI1 or GST‐AHG3, and 3 μl immunoprecipitated product in a reaction buffer containing 50 mM Tris‐HCl (pH 7.5), 10 mM MgCl2, and 10 mM ATP. After 2 h of incubation at 30°C, the reactions were immunoblotted with anti‐GST antibody and anti‐HA antibody.

In vivo ubiquitination analysis

In vivo ubiquitination assays were performed as described previously (Zhang et al., 2013). The coding sequences of ABI1 with an MYC tag (ABI1‐MYC) and AHG3 with an MYC tag (AHG3‐MYC) were cloned into the pCAMBIA1300 vector under the control of the 35S promoter. Five‐week‐old N. benthamiana leaves were used for the transient expression assay. Three days after the first infiltration, a second infiltration was carried out with infiltration buffer with or without 100 μM MG132 and then cultured for 12 h before sample harvesting. Three grams of leaf tissue from each sample was homogenized in a protein extraction buffer as described previously (Zhang et al., 2013). The supernatant (with the equal volume and the equal amount of proteins) was incubated with 15 μl anti‐MYC antibody‐conjugated agarose (Chromotek) for 3 h at 4°C. The immunoprecipitated products were washed briefly with extraction buffer three times at 4°C, and then immunoblotted with anti‐MYC antibody and anti‐Ub antibody. For the experiments investigating the effects of ABA on the ubiquitination of ABI1 and AHG3, the first infiltration was followed by dark treatment for 3 d and then the second infiltration with the buffer containing 100 μM MG132 with or without 30 μM ABA. The samples were then cultured for 12 h before harvest. ACTIN protein was used as a loading control.

Protein extraction and Western blot analysis

Transgenic seedlings with the WT background (35S:ABI1‐MYC or 35S:AHG3‐MYC) and cop1‐4 (35S:ABI1‐MYC or 35S:AHG3‐MYC) were cultured under normal growth conditions for 5 d and subsequently transferred to media containing 100 μM CHX with or without 10 μM ABA, or 100 μM CHX plus 50 μM MG132 with or without 10 μM ABA for 3 h. Total proteins were extracted and quantified using the Bradford assay. The same amount of proteins from each sample were separated using sodium dodecyl sulphate‐polyacrylamide gel electrophoresis (SDS‐PAGE). Western blotting was performed using anti‐MYC antibodies. Actin was used as the loading control.

Thermal imaging

Thermal imaging was performed as described previously (Dong et al., 2018). Wild‐type, cop1‐4, abi1‐3, ahg3‐1, cop1‐4abi1‐3, cop1‐4ahg3‐1 and cop1‐4abi1‐3ahg3‐1 seedlings grown in normal medium for 4 d were transplanted into soil with sufficient moisture. Seedlings were not watered and the soils were naturally dried. After c. 20 d, thermal images were acquired within the growth chamber using a ThermaCAMSC1000 infrared camera (FLIR System, Danderyd, Sweden). Meanwhile, the sixth leaf of each genotype was selected for far‐infrared photography. Images were analyzed using the image‐analysis program irwin reporter (v.5.31).

Phosphatase assay

The phosphatase activity was determined by using the Serine/Threonine Phosphatase Assay System (Promega) according to the manufacturer’s instructions. Briefly, 4‐wk‐old rosette leaves of WT, cop1‐4, cop1‐4abi1‐3ahg3‐1, cop1‐4abi1‐3, cop1‐4ahg3‐1 and hexpyl were harvested and incubated in stomatal solution with or without 10 μM ABA for 1 h under light. Total proteins were extracted with the buffer containing 100 mM Hepes (pH 7.8), 5 mM EDTA, 5 mM EGTA, 10 mM sodium orthovanadate (Na3VO4), 10 mM sodium fluoride (NaF), 0.5% NP‐40, 5 mM okadaic acid, 50 ng μl−1 PPase‐1 inhibitor‐1, and 1% protease inhibitor cocktail and filtered to remove inorganic phosphate (Pi) ions using Sephadex® G‐25 resin. Total proteins were quantified using the Bradford method. One‐hundred micrograms of protein was used to perform the phosphatase assay.

Detection of phosphorylation level of OST1

To detect the phosphorylated OST1 protein in vivo, 4‐wk‐old rosette leaves of WT, cop1‐4, cop1‐4abi1‐3ahg3‐1, abi1‐3, ahg3‐1, hexpyl and ost1 were harvested and incubated in stomatal solution with or without 100 μM ABA for 1 h under light. Proteins were extracted from the seedlings using the extraction buffer described in the subsection ‘Co‐immunoprecipitation (Co‐IP) assay in Arabidopsis protoplasts’, above. After proteins were transferred to a PVDF membrane, total OST1 protein was detected using anti‐OST1 antibodies (Agrisera, Vannas, Sweden) and served as loading control, while phosphorylated OST1 was detected with an anti‐pOST1 specific antibody (kindly provided by Yang Zhao at the Shanghai Center for Plant Stress Biology).

Results

The cop1 mutants show defects in ABA regulated stomatal movement

Previous studies demonstrated that COP1 constitutively inhibits stomatal opening in both darkness and light, and that COP1 is involved in ABA‐mediated microtubule depolymerization and stomatal closure (Mao et al., 2005; Khanna et al., 2014). In this study, we also observed differences between the cop1 mutants and WT in stomatal responses to ABA (Fig. 1a,b). Treatment of WT, cop1‐4 and cop1‐6 mutants (McNellis et al., 1996) and the cop1 functional complementation line GUS‐COP1 (Osterlund & Deng, 1998) with white light at an intensity of 150 µmol m−2 s−1 intensity for 2.5 h induced full stomatal opening in all genotypes (Fig. 1a). Nevertheless, both cop1 mutants showed statistically significant increases in stomatal aperture compared to WT plants (Fig. 1b). When plants were treated with ABA (1 μM for 2 h), the cop1 mutant lines showed strongly impaired ABA‐induced stomatal closure compared to WT (Fig. 1a,b), with width : length ratios of 0.573 ± 0.05, 0.584 ± 0.056 and 0.307 ± 0.051 in cop1‐4, cop1‐6 and WT, respectively (Fig. 1b). The complementation line GUS‐COP1 phenocopied WT plants in all treatments (Fig. 1a,b). These results indicate that the cop1 mutants are hyposensitive to ABA‐mediated stomatal responses under light. In addition, the ABA‐hyposensitivity of the cop1 mutant was also observed in root growth and biomass measures at the seedling stage (Fig. S1).

Fig. 1.

Fig. 1

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Abscisic acid (ABA) induced stomatal responses of wild‐type (WT),cop1 mutants andGUS‐COP1complementation plants in light and dark conditions. (a) Representative images showing stomatal apertures of Arabidopsis. Bar, 10 μm. (b) Quantification of stomatal apertures. Data are mean values ± SD; n = c.150 stomata for three independent experiments. *, P < 0.05; **, P < 0.01; Student's t‐test.

To determine whether the ABA hyposensitivity of cop1 mutants is light‐dependent, we measured stomatal responses to ABA for all genotypes in the dark and observed a strong reduction in stomata aperture in WT and GUS‐COP1 plants. By contrast, the stomatal response of cop1 mutants to ABA was again significantly impaired (Fig. 1a,b). Together, our results indicate that COP1 plays an important role in ABA‐induced stomatal closure, which seems to be independent of COP1‐mediated light signaling.

COP1 interacts with the ABA signaling components ABI1, ABI2, HAB1, HAB2 and AHG3

To study the nature of COP1 involvement in ABA‐regulated stomatal closure, yeast two‐hybrid (Y2H) assays were used to test the interaction between COP1 and known regulatory components of the ABA signaling pathway controlling stomatal movement (Table S2). Strong interactions were detected between COP1 and several PP2Cs with critical roles in ABA signaling, including ABA insensitive 1 (ABI1), ABA insensitive 2 (ABI2), Hypersensitive to ABA1 (HAB1), Homology to ABI2 (HAB2) and ABA‐hypersensitive germination 3 (AHG3) (Fig. S2a). We also verified that none of the nine PP2Cs examined here exhibited self‐activation activity in yeast (Fig. S2b).

The COP1‐PP2C interactions were also tested using BiFC assays in N. benthamiana leaves. Green fluorescence signal was visible in N. benthamiana leaves co‐expressing cGFP‐COP1 and nGFP‐PP2Cs, while no signal was detected in those co‐expressing cGFP‐COP1 and nGFP‐PP2C5 control (Fig. 2a). Co‐expression of the nuclear fluorescence marker H2B‐mCherry revealed that the interactions of COP1 with ABI1, ABI2, HAB1 and HAB2 occurred in both cytoplasm and nucleus, whereas the interaction between COP1 and AHG3 was restricted only to the nucleus (Figs 2a, S3a). The punctuated fluorescence showing the interactions between COP1 and these PP2Cs is similar to the interaction of COP1 with HY5 (Fig. S3a), which is consistent with the previously reported localization patterns of COP1 in plant cells (Liu et al., 2008; Park et al., 2017; Swain et al., 2017). Consistent with the Y2H results, AHG1, HAI1, HAI2 and HAI3 showed no interactions with COP1 in N. benthamiana leaves (Fig. S3b). However, no direct interactions were found between COP1 and ABA receptors using Y2H and BiFC (Fig. S4)

Fig. 2.

Fig. 2

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COP1 interacts with multiple PP2Cs in vivo. (a) Confocal images showing the bimolecular fluorescence complementation (BiFC) assay for the interactions of COP1 with PP2Cs. The nGFP‐PP2Cs and cGFP‐COP1 were co‐expressed in tobacco (Nicotiana benthamiana) leaves. nGFP‐PP2C5 and cGFP‐COP1 were co‐expressed in N. benthamiana as a negative control. H2B‐mCherry was used as a nuclear localization marker. Green fluorescence indicates interaction between the proteins. Red fluorescence shows the location of the nucleus. Bar, 50 μm. (b–f) Co‐IP assays. A CFP‐COP1 fusion protein was co‐expressed with the respective PP2C‐MYC fusions in Arabidopsis protoplasts. Total protein extracts from transformed protoplasts were immunoprecipitated using anti‐GFP antibody‐conjugated agarose and detected by immunoblot using anti‐MYC antibody and anti‐GFP antibody.

To further verify the interactions in vivo, we transiently expressed CFP‐COP1 and/or PP2Cs‐MYC (ABI1, ABI2, HAB1, HAB2 and AHG3) fusion proteins in Arabidopsis protoplasts and performed co‐immunoprecipitation (Co‐IP) analysis. The results showed that CFP‐COP1 co‐precipitated with ABI1‐MYC, ABI2‐MYC, HAB1‐MYC, HAB2‐MYC and AHG3‐MYC (Fig. 2b–f). Together, our results indicate that COP1 directly interacts with multiple clade A PP2Cs (ABI1, ABI2, HAB1, HAB2 and AHG3) that play critical roles in ABA signaling.

COP1 ubiquitinates ABI1 and AHG3

Since COP1 interacts with ABI1, ABI2, HAB1, HAB2 and AHG3, we reasoned that these PP2Cs could be substrates of COP1. We chose ABI1 and AHG3 for further study as they are involved in the regulation of stomatal movement (Kuhn et al., 2006; Hua et al., 2012) and show distinct interaction patterns (Fig. 2a). We first tested whether COP1 can affect the stability of ABI1 and AHG3 proteins using a semi‐in vivo degradation assay. As shown in Fig 3a, the presence of COP1 strongly enhanced the degradation of ABI1‐MYC and AHG3‐MYC, while addition of MG132, a commonly used inhibitor of proteasome‐mediated degradation, reduced the effect of COP1. These results suggest that COP1 promotes the degradation of ABI1 and AHG3, probably through the ubiquitination pathway. We therefore performed in vitro ubiquitination analysis using GST‐ABI1 and GST‐AHG3 fusion proteins purified from Escherichia coli as substrates. In the assays, HA‐tagged ubiquitin was used as a co‐substrate and CFP as a negative control. As shown in Fig. 3(b,c), ABI1 and AHG3 were detected by both anti‐GST and anti‐HA antibodies only when E1, E2, CFP‐COP1 and HA‐tagged ubiquitin were present, indicating that both proteins were ubiquitinated by E1, E2 and E3 (CFP‐COP1) ligase. The size distribution of GST‐ABI1 and GST‐AHG3 detected by both anti‐GST and anti‐HA antibodies indicates polyubiquitination of ABI1 and AHG3 by COP1. Polyubiquitinated COP1 was also detected (Fig. 3b,c), which is consistent with a previous report showing that the E3 ligase COP1 can be self‐ubiquitinated in such reactions (Seo et al., 2003).

Fig. 3.

Fig. 3

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COP1 promotes ubiquitination and degradation of ABI1 and AHG3. (a) Semi‐in vivo protein degradation assay showing increased degradation of ABI1 and AHG3 in the presence of COP1. Protein extracts containing ABI1‐MYC, AHG3‐MYC or CFP‐COP1 from transiently transformed tobacco (Nicotiana benthamiana) leaves were used in this assay. Protein abundance was detected by immunoblotting. Actin was used as a loading control. (b, c) In vitro ubiquitination assay showing COP1‐mediated ubiquitination of ABI1 (b) and AHG3 (c). The recombinant GST‐ABI1/GST‐AHG3 fusion proteins were expressed in Escherichia coli and purified. The protein extracts containing CFP‐COP1 or CFP were purified from N. benthamiana leaves transiently expressing each protein. E1 (UBE1), E2 (UBCh5b), and HA‐tagged ubiquitin (Ub) were purchased from commercial companies. ABI1, AHG3 and COP1 ubiquitination was analyzed in immunoblots using anti‐GST and anti‐HA antibodies. (d, e) In vivo ubiquitination assays showing the ubiquitination of ABI1 (d) and AHG3 (e) by COP1. ABI1‐MYC, AHG3‐MYC and CFP‐COP1 proteins were transiently expressed in N. benthamiana leaves and immunoprecipitated using anti‐MYC antibody‐conjugated agarose. The gel blots were visualized using anti‐MYC and anti‐Ub antibodies. Actin was used as a loading control.

To further verify the ubiquitination of ABI1 and AHG3 by COP1, we co‐expressed CFP‐COP1 with ABI1‐MYC or AHG3‐MYC in tobacco leaves and examined the ubiquitin status of ABI1 and AHG3 in the presence or absence of MG132. After co‐expression, total leaf proteins were immunoprecipitated using anti‐MYC antibodies, and the precipitated proteins were detected by anti‐MYC and anti‐Ub antibodies. In the absence of MG132 treatment, ABI1 protein levels in the tobacco leaves co‐expressing CFP‐COP1 and ABI1‐MYC were lower than those in the tobacco leaves expressing ABI1‐MYC alone (Fig. 3d). Similarly, co‐expression of CFP‐COP1 and AHG3‐MYC produced lower AHG3 levels than expression of AHG3‐MYC alone (Fig. 3e). These results suggest that COP1 mediates ABI1 and AHG3 degradation in plant cells. Interestingly, in the presence of MG132, tobacco leaves co‐expressing CFP‐COP1 and either ABI1‐MYC or AHG3‐MYC accumulated more polyubiquitinated ABI1‐MYC or AHG3‐MYC than leaves expressing ABI1‐MYC alone or AHG3‐MYC alone (Fig. 3d,e), likely due to inhibition of degradation of ubiquitinated proteins caused by MG132.

ABA promotes the degradation of ABI1 and AHG3 by COP1

Recent studies have revealed that ABA promotes the degradation of PP2Cs by the proteasome‐mediated pathway (Kong et al., 2015; Wu et al., 2016). Since our results show that COP1 can ubiquitinate ABI1 and AHG3, we tested the effect of ABA on the COP1‐mediated degradation of these PP2Cs. We first determined the impact of ABA on the interaction between COP1 and either ABI1 or AHG3. Co‐IP results showed that ABA treatment increased the amount of ABI1‐MYC and AHG3‐MYC proteins co‐precipitating with COP1. In addition, the amount of ABI1‐MYC and AHG3‐MYC proteins co‐precipitated with COP1 increased with increasing ABA concentrations (Fig. 4a,b). Fluorescence intensities of the protoplasts co‐expressing cLuc‐COP1 and nLuc‐ABI1, or cLuc‐COP1 and nLuc‐AHG3 were higher than those of the protoplasts co‐expressing nLuc‐ABI1 and cLuc‐Empty, nLuc‐AHG3 and cLuc‐Empty, or nLuc‐Empty and cLuc‐COP1 as indicated by split luciferase complementation assay (Fig. S5). Moreover, fluorescence intensities of the protoplasts co‐expressing cLuc‐COP1 and nLuc‐ABI1, or cLuc‐COP1 and nLuc‐AHG3 were further increased by treatment with 10 μM ABA (Fig. S5). These results suggest that ABA enhances the association between COP1 and these two PP2Cs.

Fig. 4.

Fig. 4

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Abscisic acid (ABA) promotes the degradation of PP2Cs by COP1 in vivo. (a, b) Immunoblot showing enhanced interaction between COP1 and either ABI1 (a) or AHG3 (b) by ABA. 35S:CFP‐COP1 and either 35S:ABI1‐MYC or 35S:AHG3‐MYC were co‐expressed in Arabidopsis protoplasts in the presence of different concentrations of ABA (− represents 0 μM, + represents 1 μM, ++ represents 10 μM). The total protein fraction was purified using anti‐GFP antibody‐conjugated agarose and analyzed by immunoblotting using anti‐MYC and anti‐GFP antibodies. (c) In vivo ubiquitination assay showing enhanced ubiquitination of ABI1 and AHG3 by COP1. Protein extracts from tobacco (Nicotiana benthamiana) leaves transiently co‐expressing CFP‐COP1 and either ABI1‐MYCor AHG3‐MYC, with or without 30 μM ABA treatment, were purified with anti‐MYC antibody‐conjugated agarose. The immunoprecipitated proteins were analyzed by immunoblotting using anti‐MYC and anti‐Ub antibodies. Actin was used as a loading control. (d–f) Protein stability assay showing increased stability of PP2Cs in cop1‐4. Seedlings of 35S:ABI1‐MYC and 35S:AHG3‐MYCtransgenic plants in the wild‐type (WT) and cop1‐4 backgrounds were treated with 100 μM CHX (−ABA/+ABA) or 100 μM CHX plus 50 μM MG132 (−ABA/+ABA) and the total protein extracts were analyzed by Western blotting with anti‐MYC and anti‐actin antibodies. Actin was used as a loading control. (e, f) Quantitative analysis of the signal intensity shown in panel (d). Data are mean values ± SE of three replicates. Different letters indicate groupings of statistical differences at P < 0.05 (one‐way ANOVA analysis).

We then determined the effect of ABA on COP1‐mediated ubiquitination of PP2Cs in vivo. CFP‐COP1 and either ABI1‐MYC or AHG3‐MYC were co‐expressed in tobacco leaves in the presence or absence of ABA, and anti‐MYC was used to immunoprecipitate ABI1‐MYC and AHG3‐MYC. Western blotting of the immunoprecipitated proteins showed that ABA treatment increased the polyubiquitination levels of ABI1‐MYC and AHG3‐MYC (Fig. 4c). These results indicate that ABA promotes ubiquitination of ABI1 and AHG3 by COP1 in plant tissues.

To study whether ABA regulates ABI1 and AHG3 protein stability via COP1 in Arabidopsis, we generated transgenic plants constitutively expressing either ABI1‐MYC or AHG3‐MYC fusion proteins (35S:ABI1‐MYC or 35S:AHG3‐MYC), and introduced the transgenes into the cop1‐4 mutant background by genetic crossing. In the absence of MG132, the cop1‐4 mutant accumulated more ABI1‐MYC and AHG3‐MYC than WT (Fig. 4d–f). Abscisic acid‐promoted degradation of ABI1‐MYC and AHG3‐MYC in the cop1‐4 background was less pronounced than in WT plants, and ABA treatment resulted in a decrease of ABI1‐MYC protein by 38% in WT and by 16% in cop1‐4. Similarly, AHG3‐MYC protein level was decreased by 42% in WT and by 23% in cop1‐4 after ABA treatment (Fig. 4d–f). However, when plants were treated with 50 μM MG132, which inhibits proteasome‐mediated protein degradation, there were no statistically significant differences in the ABI1‐MYC and AHG3‐MYC levels between WT and cop1‐4 backgrounds (Fig. 4e,f). These results indicate that COP1 mediates the degradation of ABI1 and AHG3 through the 26S proteasome pathway, and ABA‐enhanced ABI1 and AHG3 degradation is partly mediated by COP1 in Arabidopsis.

COP1‐mediated ubiquitination of ABI1 and AHG3 in ABA signaling

Transient co‐expression of COP1 with ABI1 or AHG3 in the hexpyl ABA receptor mutant was used to determine whether the ABA‐promoted degradation of ABI1 and AHG3 by COP1 is dependent on PYL ABA receptors . Wild‐type and hexpyl mutant protoplasts co‐transformed with equal amounts of Pro35S:CFP‐COP1, Pro35S:ABI1‐MYC or Pro35S:CFP‐COP1, Pro35S:AHG3‐MYC plasmids, respectively, were used to detect the abundance of ABI1‐MYC and AHG3‐MYC proteins (Fig. 5a). Quantitative measurements showed that the degradation rates of ABI1‐MYC and AHG3‐MYC were markedly reduced in the hexpyl mutant compared to WT with or without ABA treatment, and the ABA‐enhanced degradation of these PP2Cs in WT was also alleviated in the hexpyl mutant (Fig. 5b). Treatment with MG132 significantly reduced the degradation rate of ABI1‐MYC and AHG3‐MYC in both the WT and hexpyl mutant, suggesting that the degradation is dependent on the 26S proteasome pathway. Together, these results indicate that the ABA‐promoted degradation of ABI1 and AHG3 via COP1 is partly dependent on PYLs.

Fig. 5.

Fig. 5

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COP1 mediates the ubiquitination of ABI1 and AHG3, which are involved in the modulation of abscisic acid (ABA) signaling. (a) Semi‐in vivo degradation analysis of ABI1‐MYC and AHG3‐MYC in WT and hexpyl backgrounds by co‐expression with CFP‐COP1. After incubation for 16 h, the Arabidopsis protoplasts were treated with or without ABA, or with ABA + MG132. An anti‐MYC antibody was used to detect the ABI1‐MYC protein, and actin was used as a loading control. Values below the images are the contents of ABI1‐MYC or AHG3‐MYC relative to wild‐type (WT) at each time point following the initiation of the experiment (0h). (b) Quantitative analysis of the signal intensity of ABI1‐MYC and AHG3‐MYC, respectively. (c) Phosphatase activity in WT, cop1‐4, cop1‐4abi1‐3ahg3‐1, cop1‐4abi1‐3, cop1‐4ahg3‐1 and hexpyl plants in the absence or presence of 10 μM ABA. Total PP2C activity is represented by the phosphatase activity assay, in which 5 mM okadaic acid was added to inhibit the activity of PPP family Ser/Thr‐specific phosphoprotein phosphatases (e.g. PP1 and PP2A). Error bars represent SD (n = 6). Different letters indicate groupings of statistical differences at P < 0.05 (one‐way ANOVA analysis). (d) Detection of phosphorylated OST1 in various genotype materials. Four‐week‐old rosette leaves of WT, cop1‐4, cop1‐4abi1‐3ahg3‐1, abi1‐3, ahg3‐1, hexpyl and ost1 were harvested and incubated in stomatal solution with or without 100 μM ABA for 1 h under light. Total protein was extracted and total OST1 was used as a loading control. Detection of OST1 was performed using an anti‐OST1 antibody from Agrisera, and phosphorylated OST1 was detected using an anti‐pOST1 antibody.

We further examined the effect of COP1 on ABI1 and AHG3 enzyme activity. In the phosphatase activity assay, free phosphate was hardly detected for GST, MBP or MBP‐COP1, while a large amount of free phosphate was released by the dephosphorylation activity of GST‐ABI1 or GST‐AHG31, and the content of free phosphate produced by ABI1 or AHG3 was significantly reduced when MBP‐COP1 was added (Fig. S6). This result indicates that the presence of COP1 impairs the phosphatase activity of ABI1 and AHG3. To consolidate the role of COP1 in modulating ABA signaling through PP2Cs and the downstream components, we generated cop1‐4abi1‐3, cop1‐4ahg3‐1 and cop1‐4abi1‐3ahg3‐1 mutants by genetic crossing (Fig. S7a–d) and measured the phosphatase activity in WT, cop1‐4, cop1‐4abi1‐3ahg3‐1, cop1‐4abi1‐3, cop1‐4ahg3‐1, and hexpyl. The hexpyl mutant showed the highest phosphatase activity among all genotypes, and the cop1‐4 mutant had significantly higher phosphatase activity than WT with or without ABA treatment (Fig. 5c). Abscisic acid treatment reduced the phosphatase activity in all tested genotypes, and the abi1‐3 and ahg3‐1 mutations reduced the phosphatase activity of the cop1‐4 mutant (Fig. 5c). These results further support the idea that COP1 mediates the degradation of ABI1 and AHG3, thus affecting their phosphatase activity.

We subsequently detected the phosphorylation levels of OST1 in the leaves of WT, cop1‐4, cop1‐4abi1‐3ahg3‐1, abi1‐3, ahg3‐1, hexpyl and ost1 with or without ABA treatment. Two specific antibodies have been used, and the anti‐OST1 is an antibody specific to OST1 protein, while the anti‐pOST1 is an antibody specific to phosphorylated OST1 protein. We observed that OST1 protein was present in all genotypes except ost1, while phosphorylated OST1 could only be detected after ABA treatment (Fig. 5d). Quantitative analysis showed that the level of phosphorylated OST1 was lowest in hexpyl among all genotypes and significantly reduced in cop1‐4 when compared with WT, while abi1‐3 and ahg3‐1 mutants showed higher levels of phosphorylated OST1 than WT (Fig. 5d). Although the level of phosphorylated OST1 in cop1‐4abi1‐3ahg3‐1 was lower than that in WT, the cop1‐4abi1‐3ahg3‐1 mutant had higher phosphorylated OST1 than the cop1‐4 single mutant, indicating that the abi1‐3ahg3‐1 mutations could partially restore the phosphorylation level of OST1 in cop1‐4. Together, these results indicate that COP1 modulates the PP2C activity and consequently the kinase activity of OST1 to control stomatal movement.

The abi1‐3 and ahg3‐1 mutations suppress the hyposensitivity of cop1‐4 mutants to ABA in stomatal movement

We tested whether loss‐of‐function mutations in ABI1 and AHG3 could rescue the stomatal response phenotype of the cop1 mutant to ABA under light conditions. The reduced ABA sensitivity shown by the cop1‐4 mutant was restored to the WT level in the cop1‐4abi1‐3 and cop1‐4ahg3‐1 double mutants (Fig. 6a,b), indicating that abi1‐3 and ahg3 suppress the cop1‐4 hyposensitivity to ABA‐mediated stomatal closing. Since the cop1 mutants are hyposensitive to ABA‐mediated stomatal responses in both light and dark conditions, we also examined the stomatal responses of these genotypes to ABA under dark conditions. The stomatal opening of cop1‐4 was still larger than that of other genotypes, and the stomatal opening of cop1‐4abi1‐3, cop1‐4ahg3‐1 and cop1‐4abi1‐3ahg3‐1 were larger than that of the WT but significantly smaller than that of cop1‐4 under ABA treatment in darkness (Fig. S8). This genetic evidence further supports the idea that ABI1 and AHG3 are downstream targets of COP1 and that the reduced sensitivity of cop1 to ABA‐mediated stomatal closing is due to decreased degradation of these PP2Cs.

Fig. 6.

Fig. 6

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The Arabidopsis abi1‐3 and ahg3‐1 mutations restore cop1‐4 phenotypes. (a) Representative images showing the stomatal apertures of wild‐type (WT), cop1‐4, abi1‐3, ahg3‐1, cop1‐4abi1‐3, cop1‐4ahg3‐1 and cop1‐4abi1‐3ahg3‐1 in response to 1 μM ABA. Bar, 10 μm. (b) Quantitative analysis of the stomatal apertures of the genotypes shown in (a). Data are mean values ± SD of three independent experiments. n =  c.150 stomata. Different letters indicate groupings of statistical differences at P  < 0.05 (one‐way ANOVA analysis). (c) Water loss of detached leaves of WT, cop1‐4, abi1‐3, ahg3‐1, cop1‐4abi1‐3, cop1‐4ahg3‐1 and cop1‐4abi1‐3ahg3‐1. Three independent experiments were performed. The data are mean values ± SE of three replicates. **, P < 0.01; Student's t‐test. (d, e) Representative false‐color infrared images of WT, cop1‐4, abi1‐3, ahg3‐1, cop1‐4abi1‐3, cop1‐4ahg3‐1, and cop1‐4abi1‐3ahg3‐1 plants withholding water for 20 d. Bars, 1 cm. (f, g) Quantitative analysis of leaf temperature of the genotypes shown in (c, d). The leaf temperatures were calculated using irwin reporter v.5.31 software. Data are mean values ± SD (n = 30 plants for each condition; data are from c. 4000 measurements of square pixels from multiple leaves of each plant). Different letters indicate groupings of statistical differences at P < 0.05 (one‐way ANOVA analysis).

Water loss was measured in detached leaves of soil grown WT, cop1‐4, abi1‐3, cop1‐4abi1‐3, ahg3‐1, cop1‐4ahg3‐1 and cop1‐4abi1‐3ahg3‐1 mutants. As expected, cop1‐4 mutants showed higher water loss than WT (Fig. 6c). The increased water loss in cop1‐4 was only partially recovered in cop1‐4abi1‐3 and cop1‐4abi1‐3ahg3‐1 mutants (Fig. 6c, measurement points starting 3 h after detachment), indicating the involvement of additional PP2Cs in the stomatal response. Stomatal density measurements showed that the cop1‐4 single mutant clearly had higher stomatal density than WT, abi1‐3 and ahg3, while the cop1‐4, cop1‐4abi1‐3, cop1‐4ahg3‐1 and cop1‐4abi1‐3ahg3‐1 mutants exhibited similar stomatal densities (Fig. S9). This result suggests that both larger aperture and higher density of stomata in cop1‐4 may contribute to higher water loss compared with WT, while the differences in water loss among the cop1‐4, cop1‐4abi1‐3, cop1‐4ahg3‐1 and cop1‐4abi1‐3ahg3‐1 mutants result merely from the differences in stomatal aperture but not stomatal density.

We also measured the leaf temperatures of these genotypes under drought conditions. The cop1‐4 mutants showed statistically significant lower temperatures than WT, while the cop1‐4abi1‐3, cop1‐4ahg3‐1 and cop1‐4abi1‐3ahg3‐1 mutants partially rescued the decreases observed in cop1‐4 temperatures (Fig. 6d–g). In order to eliminate the influence of plant size on the temperature experiments, we selected the sixth rosette leaf, which is not shaded by others, to observe the leaf temperature, and the leaf temperature results with detached leaves resembled the whole plant measurements (Fig. S10). Together, our results strongly indicate that ABI1 and AHG3 are the direct downstream targets of COP1 in the ABA‐mediated control of stomatal movement.

Discussion

The COP1 protein is highly conserved in plants and mammals, and its role in plant photomorphogenesis has been extensively studied (Wang et al., 1999; Lau & Deng, 2012). COP1 has also been implicated in stomatal movement, with the cop1 mutant exhibiting insensitivity of stomatal closing in response to darkness (Khanna et al., 2014). Our findings also prove that ABA‐induced stomatal closure in cop1 mutants is greatly reduced compared to WT plants, suggesting an important role for COP1 in ABA signaling. However, the molecular basis for the role of COP1 in the ABA control of stomatal movement is still unknown. In this study, we show that COP1 interacts with several clade A PP2Cs, resulting in their ubiquitination and degradation, which is enhanced by ABA, to potentiate the ABA‐triggered stomatal closure.

This work provides several lines of evidence supporting a key role for COP1 in the ABA‐triggered stomatal closure. First, ABA‐induced stomatal closure in cop1 mutants is severely reduced compared to wild‐type plants. The ABA hyposensitivity in cop1‐4 can be partially recovered in cop1‐4abi1‐3, cop1‐4ahg3‐1 and cop1‐4abi1‐3ahg3‐1 mutants, implying that ABI1 and AHG3 are downstream targets of COP1 in regulating stomatal movement. Second, our in vitro and in vivo data show the interaction between COP1 and several clade A PP2Cs, including ABI1 and AHG3. Third, we show that COP1 can ubiquitinate ABI1 and AHG3, promoting their degradation. Importantly, ABA enhances the association of COP1 with either ABI1 or AHG3, and promotes the ubiquitination and degradation of both PP2Cs. The ABA‐induced reduction of these PP2Cs by COP1 promotes stomatal closure in response to stress conditions. In accordance, cop1 mutants do not efficiently reduce PP2C levels in response to ABA and, as a consequence, fail to accumulate high levels of phosphorylated OST1, thus inhibiting stomatal closure.

The finding that COP1 acts as an upstream positive regulator in ABA signaling raises the question of how the COP1‐clade A PP2C regulatory module fits into the ABA signaling pathway. We propose a model in which COP1 recruits and degrades a subset of the clade A PP2C co‐receptors in the ABA‐PYR/PYL/RCAR‐PP2C‐SnRK2 core signaling pathway (Fig. 7) promoting stomatal closure. Our results showing that cop1 mutants show reduced, but not completely abolished, sensitivity to ABA‐triggered stomatal closure support this model. The existence of several ABA‐mediated signaling mechanisms provides redundancy, and amplifies the speed and magnitude of the stress response. The evolution of multiple mechanisms to regulate ABA co‐receptors also reflects the importance and the complex dynamics controlling the regulation of stomatal movement.

Fig. 7.

Fig. 7

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A simplified model for the role of COP1 in abscisic acid (ABA)‐regulated stomatal closure in Arabidopsis. The clade A PP2Cs act as negative regulators of stomatal closure mainly by inhibiting the activity of OST1. Recruitment of the clade A PP2Cs by the ABA‐PYR/PYL/RCAR receptor pathway leads to de‐repression of OST1 and stomatal closure (right‐hand pathway in the model schematic depicted in the figure). COP1‐mediated ubiquitination and subsequent degradation of the clade A PP2Cs increases free OST1, leading to stomatal closure (left‐hand pathway). Abscisic acid can potentiate the effect of the COP1‐PP2Cs regulatory module. The ABA‐promoted degradation of ABI1 and AHG3 by COP1 is partly dependent on PYLs.

COP1 is likely to play a role in the dynamic modulation and homeostasis of clade A PP2C co‐receptors in the cell, even in the absence of stress, as it does not require the presence of ABA for its interaction with ABI1 and AHG3 and their subsequent ubiquitination. Nevertheless, ABA greatly increases the association of COP1 with these two substrates, and enhances the ubiquitination of ABI1 and AHG3 by COP1, linking it to the stress response. Indeed, previous studies have shown that the expression of COP1 is induced by multiple stresses, including drought, salt and cold (Moazzam‐Jazi et al., 2018). The increased COP1 levels in response to stress could incite increased COP1‐mediated degradation of some of the clade A PP2Cs promoting stomatal closure. Dynamic changes in PP2C abundance could be important for adaptation of plants to changing environmental conditions by timely and efficiently modulating stomatal behavior.

Although we have proven the role of COP1 in the control of ABI1 and AHG3 cellular levels, our protein interaction studies show strong interaction with other PP2C co‐receptors such as ABI2, HAB1 and HAB2, which suggests that COP1 might also be involved in their regulation. Indeed, the fact that the cop1‐4abi1‐3, cop1‐4ahg3‐1 and cop1‐4abi1‐3ahg3‐1 mutants can only partially rescue the cop1‐4 phenotype indicates the involvement of additional PP2Cs in this process, suggesting the involvement of other PP2Cs in regulating COP1‐related stomatal movement.

According to sequence alignment and phylogenetic analysis, clade A PP2Cs are divided into two groups, with ABI1, ABI2, HAB1 and HAB2 in one and AHG1, AHG3, HAI1, HAI2 and HAI3 in the other (Schweighofer et al., 2004; Antoni et al., 2012). The interaction of COP1 with only five clade A PP2Cs suggests that the interaction is selective. Similar selectivity has been reported for several other clade A PP2C‐interacting proteins. For example, the uncharacterized protein EAR1 (Enhancer of ABA co‐receptor 1) negatively regulates ABA signaling by interacting with six PP2C members, but not HAI (Wang et al., 2018). The E3 ligase PIR1.2 (RING finger protein1.2) interacts strongly with PP2CA and AHG1, and weakly with HAI1 and HAI3, but not with ABI/HAB PP2Cs (Baek et al., 2019). The E3 ligases RGLG (RING DOMAIN LIGASE) 1 and 5 modulate ABA signaling by controlling the turnover of AHG3, ABI2, and HAB2 (Wu et al., 2016). In addition, DOG1 (Delay of germination1), RH8 (RNA helicase‐like8), BPM (BTB/POZ AND MATH DOMAIN protein) 3 and 5 all display selective interactions with members of the clade A PP2Cs (Née et al., 2017; Baek et al., 2018; Julian et al., 2019).

On the other hand, functional divergence may explain the interaction of COP1 with AHG3 but not AHG1. Although both AHG1 and AHG3 function in ABA signaling during seed development and germination, AHG3 also plays a role in stomatal movement in response to ABA, while AHG1 is only expressed in seeds, and mutations in AHG1 show no phenotypes in adult plants (Nishimura et al., 2007; Antoni et al., 2012). Similarly, the HAI group PP2Cs are functionally distinct from other members. HAI PP2C genes are strongly induced by stress or ABA, and HAI PP2Cs participate in drought and osmotic stress response mainly by negatively regulating the accumulation of osmolytes such as proline and betaine in an ABA‐independent manner. In addition, the hai double and triple mutants show ABA‐insensitive seed germination, which is different from other clade A PP2C members (Bhaskara et al., 2012). The distinct functions of HAI from other clade A PP2Cs could explain the selective interaction of COP1 with some of the members but not HAIs. Nevertheless, detailed interaction analysis, including identifying the interacting motifs in COP1 and the clade A PP2Cs, will be needed to further understand the selective interaction of COP1 with the clade A PP2Cs.

Furthermore, our data also suggest that COP1 might not be the only E3 ligase involved in the degradation of clade A PP2Cs, since the ABI1 and AHG3 levels still decreased in the cop1‐4 mutant after ABA treatment. Our results are consistent with previous reports indicating that other E3 ligases, such as PUB12/13, RGLG1/5, and BPM3/5, are also involved in the regulation of the stability of PP2Cs (Kong et al., 2015; Wu et al., 2016; Julian et al., 2019).

Further work is needed to determine which other ABA‐mediated responses use the ABA‐induced COP1 ubiquitination and degradation of PP2C co‐receptors as part of the signaling mechanism. COP1 has been implicated in the modulation of cytoskeletal processes by degradation of microtubule proteins and activation of the S‐type anion channels. It will be interesting to investigate whether the COP1‐dependent degradation of ABI1 and AHG3, and possibly a whole subset of PP2C co‐receptors, may contribute to this process.

Our results indicate the involvement of COP1 in ABA‐induced stomatal closure in both light and darkness, since stomatal closure in cop1 mutants is still hyposensitive to ABA in dark conditions (Fig. 1). COP1 is a critical light signaling component, and it accumulates in the nucleus in darkness, while visible light promotes the nuclear export of COP1 (von Arnim et al., 1997; Lau & Deng, 2012; Hoecker, 2017). We show that COP1 interacts with ABI1, ABI2, HAB1 and HAB2 in both the cytoplasm and nucleus, providing a possible mechanism to explain the involvement of COP1 in stomatal movement. Overall, our data provide a mechanistic explanation positioning COP1 as a central integrator of the crosstalk between ABA and light signaling in the regulation of stomatal movement.

Author contributions

C‐PS conceived and directed the project. C‐PS, QC and LB designed all experiments. QC performed all experiments with the help of WW, QZ and KL. WW helped prepare the Arabidopsis mutant materials used in this study. C‐PS, LB, QC, and HS performed data analysis. HS, QC, LB, C‐PS, JRB and H‐QY wrote the manuscript with the assistance and approval of all authors. QC, LB and WW contributed equally to this work.

Supporting information

Fig. S1 The cop1‐4 mutant is hyposensitive to ABA in root growth.

Fig. S2 COP1 interacts with multiple PP2Cs in yeast.

Fig. S3 Bimolecular fluorescence complementation (BiFC) assays of interactions between COP1 and the indicated PP2Cs in Nicotiana benthamiana leaves.

Fig. S4 Interaction analysis of COP1 and PYLs.

Fig. S5 Split‐luciferase complementation assay of interactions between COP1 and ABI1/AHG3 in Arabidopsis protoplasts.

Fig. S6 COP1 affects the phosphatase activity of ABI1 and AHG3 in vitro.

Fig. S7 Identification of the double mutants and the triple mutant by polymerase chain reaction (PCR) and quantitative real time polymerase chain reaction (qRT‐PCR).

Fig. S8 Stomatal opening of different genotypes under dark conditions.

Fig. S9 Stomatal density in the leaves of different genotypes.

Fig. S10 The leaf temperature of detached leaves.

Methods S1 Supplementary Materials and Methods.

Table S1 Primer sequences used in the manuscript.

Table S2 Y2H screening results for COP1 and the known regulatory components of the ABA signaling pathway.

Please note: Wiley Blackwell are not responsible for the content or functionality of any Supporting Information supplied by the authors. Any queries (other than missing material) should be directed to the New Phytologist Central Office.

Click here for additional data file. (2.3MB, pdf)

Acknowledgements

We thank Professor Qi Xie (Institute of Genetics and Developmental Biology, Chinese Academy of Sciences) for valuable discussion, Professor Xiaohong Zhu (Henan University) and Yang Zhao (Shanghai Institutes for Biological Sciences, Chinese Academy of Sciences) for providing the pyr1pyl1pyl2pyl4pyl5pyl8 (hexpyl) seed and anti‐pOST1 specific antibody. This work was supported by National Natural Science Foundation of China (31970198) and Key Project of Natural Science Foundation of China (U1604233).

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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 The cop1‐4 mutant is hyposensitive to ABA in root growth.

Fig. S2 COP1 interacts with multiple PP2Cs in yeast.

Fig. S3 Bimolecular fluorescence complementation (BiFC) assays of interactions between COP1 and the indicated PP2Cs in Nicotiana benthamiana leaves.

Fig. S4 Interaction analysis of COP1 and PYLs.

Fig. S5 Split‐luciferase complementation assay of interactions between COP1 and ABI1/AHG3 in Arabidopsis protoplasts.

Fig. S6 COP1 affects the phosphatase activity of ABI1 and AHG3 in vitro.

Fig. S7 Identification of the double mutants and the triple mutant by polymerase chain reaction (PCR) and quantitative real time polymerase chain reaction (qRT‐PCR).

Fig. S8 Stomatal opening of different genotypes under dark conditions.

Fig. S9 Stomatal density in the leaves of different genotypes.

Fig. S10 The leaf temperature of detached leaves.

Methods S1 Supplementary Materials and Methods.

Table S1 Primer sequences used in the manuscript.

Table S2 Y2H screening results for COP1 and the known regulatory components of the ABA signaling pathway.

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