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. 2022 Oct 1;236(6):2061–2074. doi: 10.1111/nph.18472

Elevated CO2 induces rapid dephosphorylation of plasma membrane H+‐ATPase in guard cells

Eigo Ando 1,2,✉, Hannes Kollist 3, Kohei Fukatsu 2, Toshinori Kinoshita 2,4,✉, Ichiro Terashima 1
PMCID: PMC9828774  PMID: 36089821

Summary

  • Light induces stomatal opening, which is driven by plasma membrane (PM) H+‐ATPase in guard cells. The activation of guard‐cell PM H+‐ATPase is mediated by phosphorylation of the penultimate C‐terminal residue, threonine. The phosphorylation is induced by photosynthesis as well as blue light photoreceptor phototropin. Here, we investigated the effects of cessation of photosynthesis on the phosphorylation level of guard‐cell PM H+‐ATPase in Arabidopsis thaliana.

  • Immunodetection of guard‐cell PM H+‐ATPase, time‐resolved leaf gas‐exchange analyses and stomatal aperture measurements were carried out.

  • We found that light–dark transition of leaves induced dephosphorylation of the penultimate residue at 1 min post‐transition. Gas‐exchange analyses confirmed that the dephosphorylation is accompanied by an increase in the intercellular CO2 concentration, caused by the cessation of photosynthetic CO2 fixation. We discovered that CO2 induces guard‐cell PM H+‐ATPase dephosphorylation as well as stomatal closure. Interestingly, reverse‐genetic analyses using guard‐cell CO2 signal transduction mutants suggested that the dephosphorylation is mediated by a mechanism distinct from the established CO2 signalling pathway. Moreover, type 2C protein phosphatases D6 and D9 were required for the dephosphorylation and promoted stomatal closure upon the light–dark transition.

  • Our results indicate that CO2‐mediated dephosphorylation of guard‐cell PM H+‐ATPase underlies stomatal closure.

Keywords: Arabidopsis thaliana, CO2 signalling, dephosphorylation, guard cell, light–dark transition, photosynthesis, plasma membrane H+‐ATPase, stomatal closure

Introduction

Stomata, which are microscopic pores comprising pairs of guard cells in the plant epidermis, control leaf gas‐exchange in response to various environmental and endogenous stimuli. Light stimulates stomatal opening and promotes CO2 uptake for photosynthesis, whereas darkness, high CO2 concentrations, low air humidity, the phytohormone abscisic acid (ABA) and drought stress induce stomatal closure to prevent excess water loss via transpiration (Roelfsema et al., 2012; Inoue & Kinoshita, 2017; Jezek & Blatt, 2017; Lawson & Matthews, 2020).

Red and blue light induce stomatal opening via photosynthesis‐ and phototropin‐dependent mechanisms. Stomatal opening mediated by blue light‐activated phototropins is induced by the low fluence rate of blue light under high fluence rate of red light that saturates photosynthesis (Shimazaki et al., 2007; Inoue & Kinoshita, 2017). In guard cells, phototropins mediate phosphorylation of penultimate C‐terminal residue of plasma membrane (PM) H+‐ATPase, threonine (Thr), which recruits regulatory protein 14‐3‐3 to activate this enzyme (Kinoshita & Shimazaki, 1999, 2002; Kinoshita et al., 2001). The activated PM H+‐ATPase generates a H+ concentration gradient across the PM, which hyperpolarises the membrane potential, prompting voltage‐gated K+ channels and the H+‐coupled transporters to accumulate K+ and other anions in guard cells. The accumulation of solutes promotes osmotic water influx into the cells and therefore stomatal opening (Roelfsema et al., 2012; Inoue & Kinoshita, 2017; Jezek & Blatt, 2017). Among the 11 isoforms of PM H+‐ATPase in Arabidopsis thaliana (Arabidopsis; AHA1–AHA11), AHA1, which is the most abundant in guard cells, is responsible for blue light‐induced stomatal opening (Yamauchi et al., 2016). Recent studies revealed that clade D type 2C protein phosphatases (PP2C.Ds) regulate the phosphorylation of the penultimate residue of PM H+‐ATPase in guard cells (Wong et al., 2021; Akiyama et al., 2022). Phototropins also inhibit the release of anions through PM anion channels, which also contributes to PM hyperpolarisation (Marten et al., 2007; Hiyama et al., 2017). Triacylglycerols stored in lipid droplets in guard cells are broken down in a phototropin‐dependent manner to provide ATP for PM H+‐ATPase (McLachlan et al., 2016). Sugars and their metabolites may function as osmolytes and/or substrates for glycolysis and mitochondrial respiration during light‐induced stomatal opening (Daloso et al., 2016; Granot & Kelly, 2019). Blue light‐induced activation of PM H+‐ATPase mediates starch degradation, producing glucose in guard cells (Horrer et al., 2016; Santelia & Lunn, 2017; Flütsch et al., 2020).

Photosynthesis in mesophyll or guard‐cell chloroplasts, or both, are involved in red light‐induced stomatal opening (Mott et al., 2008; Fujita et al., 2013, 2019; Suetsugu et al., 2014). Decreases in the intercellular CO2 concentration (C i) following mesophyll photosynthesis alters the activity of PM ion channels, which facilitates ion accumulation in guard cells (Brearley et al., 1997; Roelfsema et al., 2002; Hosotani et al., 2021). Stomata also open via C i‐independent mechanisms (Messinger et al., 2006; Lawson et al., 2008; Matrosova et al., 2015). Furthermore, photosynthesis replenishes the ATP supply to PM H+‐ATPase (Tominaga et al., 2001; Suetsugu et al., 2014; Wang et al., 2014) and is a source of sucrose in guard cells (Flütsch et al., 2020). Red light induces photosynthesis‐dependent phosphorylation of the penultimate residue of PM H+‐ATPase in Arabidopsis guard cells (Ando & Kinoshita, 2018). Moreover, red‐light‐induced stomatal opening is suppressed in mutant plants lacking AHA1 (Yamauchi et al., 2016; Ando & Kinoshita, 2018; Flütsch et al., 2020). Therefore, PM H+‐ATPase activity is likely to be required for both blue and red‐light‐induced stomatal opening.

The mechanisms driving dark‐induced stomatal closure are poorly understood. Various factors affecting stomatal aperture in the dark have been identified in guard cells, including transcriptional regulation (Liang et al., 2005), the mitochondria‐mediated cytosolic ATP concentration (Goh et al., 2011), the cytoskeleton (Eisinger et al., 2012; Isner et al., 2017), photomorphogenesis (Mao et al., 2005; Khanna et al., 2014), and cytosolic pH‐dependent control of reactive oxygen species levels (Desikan et al., 2004; She et al., 2010; Ma et al., 2012). However, the early molecular processes involved in dark‐induced stomatal closure remain elusive.

When the plants are transferred to dark conditions, photosynthesis ceases and CO2 is emitted by respiration, and transiently by photorespiration (post‐illumination CO2 burst) in the leaves; therefore, dark‐induced stomatal closure may be partly mediated by CO2 (Engineer et al., 2016; Zhang et al., 2018; Chikov & Akhtyamova, 2019). Previous studies on CO2‐induced stomatal closure focused on the activation of PM anion channels in guard cells. In Arabidopsis, beta carbonic anhydrases, CARBONIC ANHYDRASE1 (CA1) and CA4 convert CO2 into bicarbonate (Hu et al., 2010; Xue et al., 2011), which inactivates the protein kinase HIGH LEAF TEMPERATURE1 (HT1; Hashimoto et al., 2006), a negative regulator of slow‐type anion channels in PM in guard cells (Tian et al., 2015; Hõrak et al., 2016; Tõldsepp et al., 2018). CONVERGENCE OF BLUE LIGHT AND CO2 1 (CBC1) and CBC2 may be activated by HT1 to suppress the activities of PM anion channels (Hiyama et al., 2017). Activation of PM anion channels promotes the release of anions from guard cells, which causes PM depolarisation and drives K+ release through outward‐rectifying voltage‐gated K+ channels in guard cells, leading to osmotic water efflux from the cells and stomatal closure (Roelfsema et al., 2012; Jezek & Blatt, 2017). Indeed, plants lacking anion channels exhibited delayed stomatal closure in response to dark conditions (Negi et al., 2008; Vahisalu et al., 2008; Jalakas et al., 2021). Moreover, CO2 may inhibit PM H+ extrusion pumps in guard cells (Edwards & Bowling, 1985), implying that dark‐induced stomatal closure involves the CO2‐mediated inhibition of PM H+‐ATPase in guard cells; however, molecular evidence for this hypothesis is lacking.

In this study, we found that light–dark transition induces the dephosphorylation of guard‐cell PM H+‐ATPase within 1 min in Arabidopsis leaves, accompanied by an increase in C i caused by the cessation of photosynthesis. Inspired by this finding and the hypothesis that CO2 may inhibit PM H+‐ATPase in guard cells (Edwards & Bowling, 1985), we conducted physiological and reverse‐genetic analyses using CO2 signal transduction mutants to characterise the dephosphorylation of PM H+‐ATPase. We also investigated the potential involvement of PP2C.Ds in rapid dephosphorylation and stomatal closure upon the light–dark transition. Our results indicate that elevated CO2 induces the dephosphorylation of guard‐cell PM H+‐ATPase, which facilitates stomatal closure.

Materials and Methods

Plant materials and growth conditions

Arabidopsis Columbia‐0 (Col‐0) was used as the wild‐type in physiological experiments. Col‐0 was also used as the control for blue light‐dependent h + ‐atpase phosphorylation‐1 (bhp‐1; Hayashi et al., 2017), ca1 ca4 (Hu et al., 2010), cbc1 cbc2 (Hiyama et al., 2017), ht1‐8D (Hõrak et al., 2016), ht1‐9 (Hiyama et al., 2017), open stomata1‐3 (ost1‐3; Yoshida et al., 2002) and pp2c.ds (Akiyama et al., 2022). Landsberg erecta (Ler) was used as the control for ost2‐1 (Merlot et al., 2007). Plants were grown in soil in a growth room or chamber. The light regime, growth temperature and relative humidity were set to a 16‐h photoperiod with a white fluorescence lamp (c. 50 μmol m−2 s−1), 20–24°C and 40–60%, respectively. Plants grown for 4–6 wk were kept in the dark overnight and fully expanded mature leaves were used for the experiments.

Light sources

Light‐emitting photodiodes (ISL‐150X150‐H4RHB; CCS) with a power supply (ISC‐201‐2; CCS) were used as the light source, except for in the experiment for Fig. 2 (please refer to later paragraphs), in which a halogen projector lamp (6423; Philips, Amsterdam, the Netherlands) with a red filter (2–61; Corning Inc., Corning, NY, USA) and power supply (MHAB‐150 W; Moritex, Saitama, Japan) was used. Photon flux densities were measured with a light analyser (LA‐105; NK System, Osaka, Japan).

Fig. 2.

Fig. 2

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Darkness triggered a decrease in the photosynthetic rate (A) and an increase in intercellular CO2 concentration (C i), followed by a decrease in stomatal conductance (g s). Arabidopsis thaliana wild‐type Col‐0 leaves were preilluminated with red light (600 μmol m−2 s−1) for 60 min and kept in the dark for 30 min. Grey and red bars represent darkness and red‐light illumination, respectively. Right panels show measurements taken 1 min after the termination of red light. Data are the mean ± SE of three leaves taken from different plants.

Treatments for immunohistochemical analyses and stomatal aperture measurements

Leaves collected from dark‐treated plants were put on Mill‐Q water (Millipore, Burlington, MA, USA) and treated as indicated. Isolated epidermal fragments were prepared from the dark‐treated leaves by blending the leaves for < 5 s twice in Milli‐Q water with a blender (7011HS; Waring Commercial) equipped with a container (MC‐1; Waring Commercial, Stamford, CT, USA) at high speed. The epidermal fragments were collected on 59‐μm nylon mesh and incubated in a buffer comprising 5 mM 2‐ethanesulfonic acid (MES)‐BTP (pH 6.5), 50 mM KCl and 0.1 mM CaCl2. For bicarbonate treatment experiments, the concentration of MES and the buffer pH were adjusted to 20 mM and pH 5.5, respectively. NaHCO3 or Na2HPO4 dissolved in Mill‐Q water was added to the buffer at concentrations of 2.5 and 1.25 mM, respectively. The final volume of water added to the buffer was 0.27% (v/v).

Immunohistochemical analyses of guard‐cell PM H+‐ATPase in leaves

The immunohistochemical analyses using leaves were performed according to the previous method with slight modifications (Ando & Kinoshita, 2018). In brief, leaves were chemically fixed with 4% (w/v) formaldehyde (prepared from paraformaldehyde, Wako, Osaka, Japan) and 0.3% (w/w) glutaraldehyde (Nacalai Tesque, Kyoto, Japan) in microtubule‐stabilising buffer (MTSB; 50 mM PIPES‐NaOH (pH 7.0), 5 mM MgSO4, 5 mM EGTA) for 1 h at room temperature in the dark. Before incubation with the fixative, the leaves were infiltrated with the same solution within 20 s after the light treatments. After washing with phosphate‐buffered saline (PBS; 137 mM NaCl, 8.1 mM Na2HPO4, 2.68 mM KCl and 1.47 mM KH2PO4), the leaves were incubated with pure methanol to remove Chl. The leaves were further incubated with pure xylene for 2 min at 37°C, ethanol for 5 min at room temperature and 50% ethanol (v/v; in PBS) for 5 min at room temperature, then washed with Milli‐Q water for 5 min twice and mounted on MAS‐coated microscope slides (Matsunami). Five times freeze–thawed cycles using liquid nitrogen were applied to the leaves. The specimens were digested with 1% (w/v) Cellulase R‐10 and 0.5% (w/v) Macerozyme R‐10 (Yakult) in PBS (pH 6.0; adjusted by HCl) for 1 h at 37°C, then leaf tissue except abaxial epidermis attached to the slide was removed. The epidermis left on the slide was permeabilised with 3% (v/v) Igepal CA‐630 (MP Biochemicals) and 10% (v/v) dimethyl sulfoxide in PBS for 30 min at room temperature. After 1‐h blocking with 3% (w/v) bovine serum albumin fraction V in PBS (Gibco; Thermo Fisher Scientific, Waltham, MA, USA), the primary antisera against the conserved catalytic domain of PM H+‐ATPase (anti‐H+‐ATPase) or its C‐terminal phosphorylated residue, threonine (anti‐pThr; Hayashi et al., 2010), was applied at a dilution of 1 : 2000 in the blocking solution for overnight at 4°C. Alexa Fluor 488‐conjugated secondary antibody (A11034; Invitrogen) was applied at a dilution of 1 : 500 in the blocking solution for 3 h at 37°C.

Immunohistochemical analyses of guard‐cell PM H+‐ATPase in epidermal fragments

The immunohistochemical analyses using epidermal fragments were performed as described previously (Hayashi et al., 2011; Ando & Kinoshita, 2018). In brief, epidermal fragments were fixed with 4% (w/v) formaldehyde with 0.1% (w/w) glutaraldehyde for 2 h at 4°C. After washing, epidermal fragments were transferred to pure methanol to remove Chl, then washed with Milli‐Q water and mounted on a cover glass coated with 0.1% (w/v) poly‐l‐lysine (Sigma‐Aldrich). The specimens were digested with 3% (w/v) Driselase (Sigma‐Aldrich) and 0.5% (w/v) Macerozyme R‐10 in PBS for 45 min at 37°C, then permeabilised with 3% (w/w) Triton X‐100 in PBS for 30 min at room temperature. Blocking and antisera application were conducted as described above.

Fluorescence microscopy

After secondary antibody application, specimens were washed and covered with 50% (v/v) glycerol. The phosphorylation levels and amounts of guard‐cell PM H+‐ATPase was estimated based on Alexa Fluor 488 fluorescence intensity using ImageJ software (National Institutes of Health; Schneider et al., 2012; Ando & Kinoshita, 2018). In brief, the difference between the mean grey value of guard cells and that of the neighbouring area excluding guard cells was calculated for each pair of guard cells. The difference between the geometric mean obtained for each antiserum and normal serum was calculated as net fluorescence intensity and expressed relative to corresponding control. Data are represented by the mean ± SD of at least three independent measurements.

Immunoblot analyses of PM H+‐ATPase in guard‐cell protoplasts (GCPs)

GCPs were enzymatically isolated as described previously (Ueno et al., 2005) with slight modifications in which 0.05% (w/v) Macerozyme R‐10 and 0.0075% (w/v) pectolyase Y‐23 (Kyowa Chemical Products; Kyowa Kasei Co. Ltd, Osaka, Japan) was used for first‐ and second‐step digestion, respectively. Isolated GCPs were kept in the dark on ice until use (at least for 1 h). GCPs in a suspension buffer comprising 20 mM MES‐KOH (pH 5.5), 10 mM KCl, 0.4 M mannitol and 1 mM CaCl2, was stirred at 300 round min−1 in a glass vessel enclosed in a water jacket filled with circulating water (24°C) during the treatments. NaHCO3 dissolved in the suspension buffer was applied to GCPs at a concentration of 10 mM. The final volume of the buffer added to the suspension was 1.09% (v/v). The GCPs' proteins were subjected to immunoblot analyses using primary antisera anti‐H+‐ATPase and anti‐pThr. 14‐3‐3 proteins were detected as a loading control using anti‐14‐3‐3 antiserum (Kinoshita & Shimazaki, 1999). The protein amounts were estimated using ImageJ software based on chemiluminescence from a reaction of horseradish peroxidase‐conjugated secondary antibody (1706515; Bio‐Rad) with the substrate (Thermo Fisher Scientific). Data are represented by the mean ± SD of three independent measurements.

Measurement of stomatal aperture

After the treatments, epidermal fragments were collected using a nylon mesh. Leaves were collected and the epidermal fragments were obtained as described above. Collected epidermal fragments were immediately microphotographed using a microscope (BX50; Olympus, Tokyo, Japan) equipped with a charge‐coupled device (CCD) camera system (DP71; Olympus). Stomatal apertures on the abaxial side fragments were measured using ImageJ software. Representative values of independent measurements were calculated as means of at least 30 stomatal apertures. Data are provided as the means of representative values obtained in at least three independent measurements with SDs.

Gas‐exchange measurements

Gas‐exchange measurements were performed using the LI‐6400 system (Li‐Cor, Lincoln, NE, USA). Air flow rate and leaf temperature were kept at 500 μmol s−1 and 23°C, respectively. Relative humidity in the leaf chamber was maintained at 40–60%. C a was set to 400 μmol mol−1 unless otherwise stated. After the measurements, the leaves were photographed and leaf areas were measured using ImageJ software. The recorded values were normalised to the true leaf area. Data are the mean ± SE of at least four separate leaves from individual plants.

To perform the combined gas‐exchange and immunohistochemical analyses, leaves were detached from the plants and immediately infiltrated with fixative. Leaf area was measured after fixation to correct the gas‐exchange measurements, as described above. The phosphorylation level and amount of guard‐cell PM H+‐ATPase are represented by the mean ± SE of four independent plants.

Estimation of stomatal density

According to a previously described method (Kang et al., 2009), leaves used for the gas‐exchange analyses were fixed with 95% (v/v) ethanol, rehydrated in 75% (v/v), 50% (v/v) and 25% (v/v) ethanol series, washed with Milli‐Q water and immersed in clearing solution (glycerol : chloral hydrate : water, 1 : 8 : 1, v/w/v). The average stomatal density of six areas of each leaf was calculated microscopically, as an estimate for the whole leaf. Data are the mean ± SE.

Statistical analyses

Means expressed as relative values were analysed by one‐sample t tests to compare with the corresponding controls, which were set to 1. The statistical significance of the difference between two independent means was assessed using Student's t‐test. For means expressed as absolute values, Dunnett's test was applied for comparison with a single control. The statistical significance of differences among means was assessed by Tukey's test. P‐values were calculated in R software using the package multcomp (Hothorn et al., 2008; R Core Team, 2021). The statistical significance of correlations was analysed using the R function cor.test. A P‐value < 0.05 was considered statistically significant.

Results

Rapid dephosphorylation of guard‐cell PM H+‐ATPase in response to the light–dark transition in leaves

Our previous immunohistochemical study using Arabidopsis leaves revealed that red light induces photosynthesis‐dependent phosphorylation of the penultimate C‐terminal residue of PM H+‐ATPase, Thr, in guard cells during red‐light‐induced stomatal opening (Ando & Kinoshita, 2018). This finding implied that the cessation of photosynthesis may affect the phosphorylation level of guard‐cell PM H+‐ATPase. To test this hypothesis, we conducted immunohistochemical analyses of Arabidopsis wild‐type Col‐0 leaves undergoing the light–dark transition using antisera against the phosphorylated Thr of PM H+‐ATPase (Hayashi et al., 2010). As the phosphorylation of PM H+‐ATPase in guard cells is mediated by both red and blue light (Inoue & Kinoshita, 2017), we first illuminated leaves with red light only (600 μmol m−2 s−1) and then subjected them to dark treatment for 1, 5, 15 and 30 min. We observed a rapid decrease in the phosphorylation of PM H+‐ATPase in guard cells, even with only a 1‐min dark treatment (Fig. 1a,b). The phosphorylation level remained low until 30 min after dark treatment (Fig. 1b). To investigate whether the decrease in phosphorylation level was caused by the change in the amount of PM H+‐ATPase itself, we conducted immunohistochemical analyses using antibodies that recognise the conserved catalytic domain of PM H+‐ATPase (Hayashi et al., 2010); there was no significant difference in the amount of PM H+‐ATPase before and after dark treatment (Fig. 1c). These results indicated that dephosphorylation of PM H+‐ATPase occurred in guard cells within 1 min of dark treatment.

Fig. 1.

Fig. 1

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Rapid dephosphorylation of guard‐cell plasma membrane (PM) H+‐ATPase in Arabidopsis thaliana wild‐type Col‐0 leaves upon light–dark transition. The phosphorylation level (a–c) and amount (d–f) of PM H+‐ATPase in guard cells were analysed by applying immunohistochemical techniques to leaves. Mature leaves harvested from dark‐treated plants were kept in the dark (Dk), illuminated with red light (RL; at 600 μmol m−2 s−1) for 30 min, kept in the dark again for the indicated time (RL→Dk), illuminated with blue light (5 μmol m−2 s−1) superimposed on red light for 2.5 min after RL (RL→RB), illuminated with red light, or kept in the dark for 1 min after RB (RB→RL, RB→Dk). (a, d) Representative fluorescence images of the phosphorylation level of PM H+‐ATPase in guard cells. Arrowheads indicate guard cells. Bar, 50 μm. (b, e) Estimation of the phosphorylation level of PM H+‐ATPase in guard cells. Asterisks indicate that the means of RL→Dk and RB→Dk are significantly lower than those of RL (b) and RL→RB (e), respectively. *, P < 0.01; **, P < 0.005; ns, not significant, P > 0.6 (one‐tailed Dunnett's test). (c, f) Estimation of the amount of PM H+‐ATPase in guard cells. The means denoted in brackets are not statistically significantly different to that of RL (c) or RL→RB (f), as indicated by the P‐values (two‐tailed Dunnett's test). Data represent the mean ± SD relative phosphorylation level and amount of PM H+‐ATPase compared with Dk, based on three (b, c) or four (e, f) independent measurements.

Next, we examined whether dark‐induced rapid dephosphorylation of PM H+‐ATPase also occurred after blue light illumination. We illuminated the leaves with weak blue light (5 μmol m−2 s−1) superimposed on the red light (600 μmol m−2 s−1) and then turned off either the blue light only or both lights and maintained those conditions for 1 min. We found that the phosphorylation level of PM H+‐ATPase was maintained for 1 min under red light following blue light treatment (Fig. 1d,e). By contrast, when both light sources were turned off, phosphorylation decreased within 1 min, with no change in the amount of PM H+‐ATPase (Fig. 1d–f); this indicates that the rapid dark‐induced dephosphorylation of PM H+‐ATPase also occurs following blue light illumination. Moreover, this rapid dephosphorylation was not observed in isolated epidermal fragments, and the dephosphorylation kinetics were comparable between red light and dark conditions (Fig. S1a). Taken together, these results indicated that guard‐cell PM H+‐ATPase is rapidly dephosphorylated during the light–dark transition in the leaves of Arabidopsis. To simplify the downstream experiments, only red light was applied in those involving the light–dark transition of leaves.

Dark‐induced stomatal closure follows cessation of photosynthesis and dephosphorylation of guard‐cell PM H+‐ATPase and requires deactivation of PM H+‐ATPase in leaves

To examine the interrelationships among the photosynthetic rate, C i and stomatal movement during the light–dark transition, we performed time‐resolved gas‐exchange analyses. As in previous studies (Engineer et al., 2016; Chikov & Akhtyamova, 2019), the photosynthetic rate decreased immediately and reached its minimum value within 1 min after the light–dark transition (Fig. 2). Similarly, the C i increased to a level slightly above the ambient CO2 concentration (C a) within 1 min of the light–dark transition. Stomatal conductance was maintained at 1 min after the light–dark transition and began to decrease c. 3 min after transfer to dark conditions. Taken together, these results suggest that the dephosphorylation of PM H+‐ATPase in guard cells is accompanied by the cessation of photosynthesis and an increase in C i, followed by stomatal closure in the leaf after the light–dark transition.

The phosphorylation of the penultimate residue of PM H+‐ATPase, Thr, is important for the activation of this enzyme through the interaction with 14‐3‐3 protein in guard cells, which creates the driving force for stomatal opening (Kinoshita & Shimazaki, 1999, 2002). To genetically validate the requirement of the deactivation of PM H+‐ATPase in dark‐induced stomatal closure, we compared ost2‐1, in which AHA1 is mutated and constitutively activated (Merlot et al., 2007) and corresponding control Ler on the stomatal closure. The light–dark transition of Ler leaves initiated decrease in the stomatal aperture within 10 min post‐transition, whereas dark‐induced stomatal closure was abolished in ost2‐1 leaves (Fig. S2). Therefore, deactivation of PM H+‐ATPase is most likely to be required for stomatal closure in response to the light–dark transition.

Elevated CO2 treatment induces the dephosphorylation of PM H+‐ATPase in guard cells

The lack of rapid dark‐induced dephosphorylation of guard‐cell PM H+‐ATPase in isolated epidermal fragments (Fig. S1a) suggested that the rapid dephosphorylation seen in the leaf was caused by the cessation of mesophyll photosynthesis upon the light–dark transition. When mesophyll photosynthesis ceases under dark conditions, the C i increases (Fig. 2; Engineer et al., 2016; Chikov & Akhtyamova, 2019), while CO2 inhibits PM H+‐ATPase in guard cells (Edwards & Bowling, 1985). This suggests that elevated C i may induce the dephosphorylation of PM H+‐ATPase in guard cells. To test this hypothesis, we combined immunohistochemical analyses with gas‐exchange measurements; leaves were treated with elevated CO2 concentrations (eCO2) in a leaf chamber (Figs 3a, S3) and immunohistochemical analyses were performed immediately. eCO2 treatment was applied for 1 or 2 min, because the C i stabilised within 2 min (Fig. S3). Without eCO2 treatment, C a was maintained at 400 μmol mol−1, and red‐light‐induced phosphorylation and dark‐induced dephosphorylation of PM H+‐ATPase in guard cells were both observed, along with the accompanying changes in the photosynthetic rate and C i (Figs 3b, S3). Under the eCO2 treatment conditions, in which C a was raised from 400 to 800 μmol mol−1 under the red light, the phosphorylation level of PM H+‐ATPase decreased within 2 min; therefore, the phosphorylation level and C i were negatively correlated under the red light (Figs 3b, S3). The amount of guard‐cell PM H+‐ATPase did not differ between treatments (Fig. 3c). These results indicated that increased C i induced the dephosphorylation of guard‐cell PM H+‐ATPase in the leaves, even in the presence of red light.

Fig. 3.

Fig. 3

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CO2 induces dephosphorylation of guard‐cell plasma membrane (PM) H+‐ATPase in Arabidopsis thaliana wild‐type Col‐0. (a–c) Elevated CO2 (eCO2 ) treatment to the leaves under red light. (a) Schematic diagram of the treatment conditions. Grey and red bars represent dark and red‐light illumination (600 μmol m−2 s−1) conditions, respectively. Green lines represent exposure to eCO2 (up to 800 μmol−1). Leaves from dark‐treated plants were treated as indicated in the leaf chamber of the gas‐exchange measurement system. (b, c) Relationships between the phosphorylation level (b) and amount (c) of guard‐cell PM H+‐ATPase estimated by immunohistochemical analyses and the intercellular CO2 concentration (C i) at the time of leaf harvesting. There was a significant negative correlation between the phosphorylation level and C i under red light (dotted line). Data represent the mean ± SE of four leaves taken from different plants. The gas‐exchange measurements recorded in this experiment are shown in Fig. S2. (d, e) Exogenous bicarbonate treatment to the isolated epidermal fragments upon the light–dark transition. (d) The phosphorylation level of guard‐cell PM H+‐ATPase estimated by immunohistochemical analyses. Isolated epidermal fragments were prepared from dark‐treated plants and kept in the dark (Dk), illuminated with red light (600 μmol m−2 s−1) for 30 min (RL), and illuminated with blue light (5 μmol m−2 s−1) superimposed onto red light for 2.5 min after RL (RL→RB), or kept in the dark for 1 min after blue light illumination (RB→Dk) with 1.25 mM Na2HPO4 , 2.5 mM NaHCO3 , or H2O. Data represent the mean ± SD relative phosphorylation levels compared with Dk from four independent measurements. Asterisks indicate that the mean is significantly lower than that of RL→RB. *, P < 0.05; ns, P > 0.1 (one‐tailed Dunnett's test). (e) Stomatal aperture analyses. The isolated epidermal fragments were kept in the dark, illuminated with blue light superimposed on red light for 2 h (RB), or kept in the dark for 10 min after RB (RB→Dk) with Na2HPO4 , NaHCO3 or H2O. Chemical concentrations and light intensity were the same as in (d). Data represent the mean ± SD of three independent measurements. Asterisks indicate that the mean is significantly lower than that of RB. *, P < 0.05; ns, P > 0.5 (one‐tailed Dunnett's test).

In the isolated epidermal fragments, the dephosphorylation of guard‐cell PM H+‐ATPase and stomatal closure occurred much more slowly (Fig. S1a,b). To further investigate whether CO2 mediates the dephosphorylation of PM H+‐ATPase in guard cells, we examined the effects of exogenous CO2 on stomatal response speeds in the epidermal fragments. We used sodium bicarbonate (NaHCO3) as an exogenous source of CO2 (Kolla et al., 2007) and investigated its effects on stomatal responses. Epidermal fragments were suspended in buffer containing 20 mM MES (pH 5.5) and preilluminated with blue light superimposed onto red light to induce the phosphorylation of guard‐cell PM H+‐ATPase (Fig. 3d, red light (RL) → red‐blue (RB)), and then transferred to dark conditions with or without NaHCO3 supplementation and incubated for 1 min. In the control conditions, in which the samples were treated with water or 1.25 mM disodium phosphate (Na2HPO4) instead of 2.5 mM NaHCO3, guard‐cell PM H+‐ATPase was not dephosphorylated 1 min after the light–dark transition; however, dephosphorylation was accelerated in samples treated with NaHCO3 (Fig. 3d). In addition, NaHCO3 induced the dephosphorylation even under blue light‐illuminated conditions, albeit with slight delay (Fig. S4). Immunoblot analyses using GCPs confirmed that NaHCO3 decreases the phosphorylation level relative to the protein amount without unequivocal protein degradation, which suggests that dephosphorylation rather than degradation of PM H+‐ATPase occurs in response to CO2 (Fig. S5). These results indicated that the rapid dephosphorylation of guard‐cell PM H+‐ATPase in leaves could be mimicked using NaHCO3. Consistent with the phosphorylation levels of PM H+‐ATPase, the exogenous NaHCO3 also enhanced dark‐induced stomatal closure; the stomatal apertures decreased within 10 min of the light–dark transition (Fig. 3e). Taken together, these results demonstrated that CO2 promotes the dephosphorylation of PM H+‐ATPase in guard cells, as well as stomatal closure, in the dark.

Carbonic anhydrases CA1 and CA4 are involved in dark‐induced dephosphorylation of PM H+‐ATPase in guard cells

Our results suggest that the molecular mechanism driving rapid dark‐induced dephosphorylation of PM H+‐ATPase may share signalling components with CO2‐induced stomatal closure (Zhang et al., 2018; Dubeaux et al., 2021). Therefore, we applied a reverse‐genetic approach. We first investigated whether the beta carbonic anhydrases CA1 and CA4, which are early signalling components in CO2‐induced stomatal closure (Hu et al., 2010), are required for dephosphorylation. We found that the phosphorylation level of guard‐cell PM H+‐ATPase was two‐fold higher in ca1 ca4 mutants compared with the wild‐type, in dark‐treated leaves (Fig. 4a, dark (Dk)). Red‐light illumination induced further phosphorylation of PM H+‐ATPase in the leaves of ca1 ca4; the PM H+‐ATPase phosphorylation level was c. 64% higher in the mutant compared with the wild‐type under RL (Fig. 4a, RL). Red light–dark transition did not induce rapid dephosphorylation of PM H+‐ATPase in ca1 ca4 leaves (Fig. 4a, RL→Dk). The amount of PM H+‐ATPase in guard cells was comparable between ca1 ca4 and the wild‐type (Fig. 4b). Furthermore, eCO2‐induced dephosphorylation was impaired in ca1 ca4 leaves (Fig. S6). These results indicated that CA1 and CA4 are required for the rapid dephosphorylation of guard‐cell PM H+‐ATPase upon the light–dark transition or C i elevation.

Fig. 4.

Fig. 4

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Lack of both CA1 and CA4 delays dark‐induced stomatal responses in Arabidopsis thaliana leaves. (a) Immunohistochemical analyses of the phosphorylation of plasma membrane (PM) H+‐ATPase in guard cells. Mature leaves harvested from dark‐treated Col‐0 and ca1 ca4 plants were kept in the dark (Dk), illuminated with red light (600 μmol m−2 s−1) for 30 min (RL), or kept in the dark for 1 min after RL (RL→Dk). Data represent mean ± SD relative phosphorylation levels compared with Col‐0 (Dk) from three independent measurements. The dagger symbol denotes that the mean is significantly higher than Col‐0 (Dk) set to 1. †, P < 0.05 (one‐tailed one‐sample t‐test). Asterisks indicate that the mean of ca1 ca4 is significantly higher than that of Col‐0. *, P < 0.05; **, P < 0.01 (one‐tailed Student's t‐test). (b) Immunohistochemical analyses of the amount of PM H+‐ATPase in guard cells. Data represent the mean ± SD relative amounts of PM H+‐ATPase compared with Col‐0 from three independent measurements. ns, mean is not significantly different from Col‐0 set to 1. P > 0.9 (two‐tailed one‐sample t‐test). (c–e) Gas‐exchange analyses in Col‐0 and ca1 ca4. Grey and red bars represent dark and red‐light illumination conditions, respectively. The light intensity applied in this experiment was the same as in (a). (c) Stomatal conductance (g s), photosynthetic rate (A) and intercellular CO2 concentration (C i). (d) g s at each time point. Asterisks indicate that the mean of ca1 ca4 is significantly higher than that of Col‐0. *, P < 0.05; ***, P < 0.001; ns, P > 0.5 (one‐tailed Student's t‐test). (e) Relative g s during the light–dark transition. g s was normalised to the value at 60 min when the light was turned off. Data represent the mean ± SE of eight (Col‐0) or seven (ca1 ca4) independent leaves from different plants.

Next, we investigated stomatal movement in ca1 ca4 by examining gas‐exchange. Although the stomatal conductance in ca1 ca4 was comparable with that of the wild‐type at the beginning of the experiment, it was higher in ca1 ca4 compared with the wild‐type following red‐light illumination (Fig. 4c,d), consistent with the higher phosphorylation level of guard‐cell PM H+‐ATPase in the mutant leaves under red‐light conditions (Fig. 4a, RL). Termination of the red‐light illumination induced a decrease in stomatal conductance in both ca1 ca4 and wild‐type leaves; however, the rate of decrease was slower in ca1 ca4 (Fig. 4c,e). Therefore, the stomatal conductance was higher in ca1 ca4 than wild‐type at 30 min after the light–dark transition (Fig. 4c,d). Photosynthesis was not impaired in ca1 ca4 (Fig. 4c). A previous study showed that ca1 ca4 exhibits increased stomatal density (Hu et al., 2010); however, we did not observe the phenotype in this study (Fig. S7a). This might be due to the conditional effects during the plant growth. Therefore, the observed differences in stomatal responses were probably not caused by differences in stomatal density. Taken together, these results indicated that CA1 and CA4 accelerated the dark‐induced dephosphorylation of PM H+‐ATPase and stomatal closure.

HT1‐mediated signal transduction is unlikely to function as a negative regulator for dark‐induced dephosphorylation of guard‐cell PM H+‐ATPase

The Raf‐like protein kinase HT1 functions as a negative regulator of CO2‐induced stomatal closure downstream of CA1 and CA4 (Tian et al., 2015; Hõrak et al., 2016; Tõldsepp et al., 2018). Thus, HT1 may function as a negative regulator of guard‐cell PM H+‐ATPase dephosphorylation. Therefore, we expected the decrease in phosphorylation levels of guard‐cell PM H+‐ATPase in the loss‐of‐function mutant. However, this phenotype was not present in a kinase‐dead mutant ht1‐9 (Fig. S8a,b). Moreover, we did not detect any increases in phosphorylation levels of guard‐cell PM H+‐ATPase in the dominant mutant ht1‐8D, in which the CO2‐medited inhibition of HT1 is impaired (Hõrak et al., 2016; Fig. S8a,b). In addition, a mutant plant lacking CBC1 and CBC2, which also belong to the Raf‐like kinases family and may be activated by HT1 to suppress CO2‐induced stomatal closure (Hiyama et al., 2017), exhibited similar phenotypes to those of ht1‐9 (Fig. S8c,d). By contrast with ht1‐9 and cbc1 cbc2 mutant plants, a mutant plant lacking BHP, another Raf‐like kinase in guard cells (Hayashi et al., 2017), exhibited decreased phosphorylation levels of guard‐cell PM H+‐ATPase regardless of the light conditions (Fig. S9). Taken together, these results suggested that the HT1‐mediated signal transduction does not function as a negative regulator of dark‐induced guard‐cell PM H+‐ATPase dephosphorylation.

OST1 does not affect the dark‐induced dephosphorylation of PM H+‐ATPase in guard cells

ABA signalling affects CO2‐induced stomatal closure, even under nonstress conditions (Xue et al., 2011; Merilo et al., 2013; Chater et al., 2015; Tian et al., 2015; Hsu et al., 2018; Dittrich et al., 2019). OST1 encodes a sucrose nonfermenting 1‐related protein kinase (SnRK) 2, SnRK2.6, a core component of ABA signalling expressed in guard cells (Mustilli et al., 2002; Yoshida et al., 2002). In leaves of the knock‐out mutant ost1‐3, neither darkness nor elevated CO2 levels induce rapid stomatal closure (Merilo et al., 2013). Therefore, a lack of OST1 might prevent the dephosphorylation of guard‐cell PM H+‐ATPase and impair stomatal closure. However, there were no detectable dephosphorylation impairments in the guard‐cell PM H+‐ATPase in ost1‐3 (Fig. S10a,b). Therefore, the dark‐induced dephosphorylation of PM H+‐ATPase in guard cells may not require OST1 activity.

PP2C.D6 and D9 are required for dark‐induced dephosphorylation of PM H+‐ATPase in guard cells

Recent studies have shown that PP2C.Ds dephosphorylate guard‐cell PM H+‐ATPase to control stomatal aperture. Among nine isogenes of PP2C.Ds, PP2C.D6 and PP2C.D9 are suggested to be major transcripts in guard cells (Wong et al., 2021; Akiyama et al., 2022). To evaluate the involvement of PP2C.D6 and/or D9 in the dark‐induced dephosphorylation of guard‐cell PM H+‐ATPase in leaves, we examined pp2c.d6 and d9 single knock‐out, and pp2c.d6/9 double knock‐out mutants. Interestingly, mutants lacking PP2C.D6 (namely pp2c.d6 and d6/9) failed to induce the rapid dephosphorylation of PM H+‐ATPase, whereas the pp2c.d9 single knock‐out mutants showed a response similar to the wild‐type upon the light–dark transition (Fig. 5a). pp2c.d6/9 leaves did not exhibit eCO2‐induced dephosphorylation of guard‐cell PM H+‐ATPase (Fig. S6). There were no differences in the amount of PM H+‐ATPase between wild‐type and mutant leaves (Fig. 5b). These results indicated that PP2C.D6 mediates the rapid dephosphorylation of guard‐cell PM H+‐ATPase upon the light–dark transition. In addition, leaves maintained in the dark exhibited different properties. Dark‐treated pp2c.d6 and wild‐type PM H+‐ATPase in guard cells exhibited comparable phosphorylation levels, whereas the phosphorylation levels in the dark‐treated pp2c.d9 and pp2c.d6/9 leaves were almost twice as high as those of the wild‐type; although, the increase in phosphorylation levels in pp2c.d9 was not statistically significant (Fig. 5a, Dk). These results suggest that both PP2C.D6 and PP2C.D9 are required to maintain PM H+‐ATPase in a dephosphorylated state after the light–dark transition.

Fig. 5.

Fig. 5

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PP2C.D6 and D9 regulate the dephosphorylation of guard‐cell plasma membrane (PM) H+‐ATPase to promote stomatal closure in Arabidopsis thaliana leaves under the light–dark transition. (a) Immunohistochemical analyses of the phosphorylation of plasma membrane (PM) H+‐ATPase in guard cells. Mature leaves harvested from dark‐treated Col‐0, pp2c.d6, d9 and d6/9 plants were kept in the dark (Dk), illuminated with red light (600 μmol m−2 s−1) for 30 min (RL), or kept in the dark for 1 min after RL (RL→Dk). Data represent mean ± SD relative phosphorylation levels compared with Col‐0 (Dk) from three independent measurements. The dagger symbol denotes that the mean is significantly higher than Col‐0 (Dk) set to 1. †, P < 0.05 (one‐tailed one‐sample t‐test). Asterisks indicate that the mean value for the mutant is significantly higher than that of Col‐0. *, P < 0.05; **, P < 0.01; ns, not significant, P > 0.4 (one‐tailed Dunnett's test). (b) Immunohistochemical analyses of the amount of PM H+‐ATPase in guard cells. Data represent the mean ± SD relative amounts of PM H+‐ATPase compared with Col‐0 from three independent measurements. ns, mean is not significantly different from Col‐0 set to 1. P > 0.9 (two‐tailed one‐sample t‐test). (c–e) Gas‐exchange analyses in Col‐0, pp2c.d6 and pp2c.d6/9. Grey and red bars represent dark and red‐light illumination conditions, respectively. The light intensity applied in this experiment was the same as in (a). (c) Stomatal conductance (g s), photosynthetic rate (A) and intercellular CO2 concentration (C i). (d) g s at each time point. Asterisks indicate that the mean value for the mutant is significantly higher than that of Col‐0. *, P < 0.05; **, P < 0.001; ns, P > 0.1 (one‐tailed Dunnett's test). (e) Relative g s during the light–dark transition. g s was normalised to the value at 60 min when the light was turned off. Data are the mean ± SE of four (Col‐0), six (pp2c.d6), or five (pp2c.d6/9) independent leaves from different plants.

Next, we performed gas‐exchange analyses to assess stomatal movement in pp2c.d6 and pp2c.d6/9. At the beginning of the measurements, stomatal conductance was higher in pp2c.d6/9 compared with the wild‐type, while the stomatal conductance of pp2c.d6 and wild‐type were comparable (Fig. 5c,d). There were no significant differences in stomatal conductance between the wild‐type and mutants under RL. In pp2c.d6/9, the dark‐induced decrease in stomatal conductance was delayed, and the stomatal conductance was higher compared with the wild‐type at 30 min after the light–dark transition (Fig. 5c–e). The decrease in stomatal conductance in pp2c.d6 was similar to that of wild‐type, although the stomatal conductance at 30 min after the light–dark transition was slightly higher in pp2c.d6 compared with the wild‐type (Fig. 5d,e). Each plant line showed similar photosynthetic rates and stomatal densities, which suggests that the observed differences in stomatal responses were not due to variations in photosynthetic capacity or stomatal density (Figs 5c, S7b). Taken together, these results indicated that PP2C.D6 and D9 control the rate of stomatal closure in leaves in the dark.

Discussion

By contrast with light‐induced stomatal opening, knowledge regarding dark‐induced stomatal closure is limited, and the early molecular mechanisms governing this process are yet to be elucidated. This study indicates that the dephosphorylation of guard‐cell PM H+‐ATPase, accompanied by an increase in C i, is an early molecular event in the course of dark‐induced stomatal closure in leaves (Figs 1, 2). We genetically validated that constitutive activation of AHA1 results in impaired dark‐induced stomatal closure in leaves, which suggests that deactivation of PM H+‐ATPase is probably required for stomatal closure upon the light–dark transition (Fig. S2). The phosphorylated penultimate residue of PM H+‐ATPase serves as a binding site for 14‐3‐3 protein, which is crucial for activation of PM H+‐ATPase in guard cells (Kinoshita & Shimazaki, 2002). Therefore, the delayed stomatal closure seen in mutant plants with dephosphorylation defects, such as ca1 ca4 and pp2c.d6/9 (Figs 4, 5), indicates that the rapid dephosphorylation of PM H+‐ATPase may facilitate deactivation of this enzyme and stomatal closure in the dark.

In this study, we found that elevated CO2 levels induce the dephosphorylation of PM H+‐ATPase in guard cells (Figs 3, S4, S5). Impairment of dark‐ or eCO2‐induced dephosphorylation of guard‐cell PM H+‐ATPase in ca1 ca4 and pp2c.d6/9 supports the hypothesis that CO2 induces the dephosphorylation (Figs 4a, 5a, S6). Although previous evidence suggests that CO2 inhibits PM H+‐ATPase in guard cells (Edwards & Bowling, 1985), the molecular mechanism behind this has not been investigated. This study suggests that CO2 inhibits PM H+‐ATPase via dephosphorylation in guard cells.

A previous mathematical modelling suggested that the conversion of CO2 into bicarbonate is delayed, not abolished, in guard cells lacking the CA activity; indeed, even ca1 ca4 exhibited stomatal closure when the plants were exposed to prolonged eCO2 conditions (Hu et al., 2015). This may be the case in prolonged dark conditions as ca1 ca4 and wild‐type plants showed comparable stomatal conductance before light illumination, at which the plants had been kept in the dark overnight before the experiments (Fig. 4c). Moreover, increased phosphorylation level of guard‐cell PM H+‐ATPase (Fig. 4a; Dk) may imply that some mechanisms counteract PM H+‐ATPase to achieve stomatal closure in the dark‐treated ca1 ca4. Such a compensatory mechanism was found in mutant plants with a defect in the slow‐type anion channel, in which elevated cytosolic Ca2+ and increased Ca2+ sensitivity of the inward‐rectifying K+ channel downregulate stomatal opening in the mutant guard cells (Laanemets et al., 2013). Further investigations would be required to elucidate the compensatory mechanisms in ca1 ca4.

The molecular pathway that drives PM H+‐ATPase dephosphorylation, which is downstream of CA1 and CA4, may differ from the CO2‐mediated regulation of PM anion channels; our data suggest that the dephosphorylation is not regulated by HT1 and CBC inhibition (Fig. S8; Zhang et al., 2018; Dubeaux et al., 2021). Nevertheless, phosphorylation levels of guard‐cell PM H+‐ATPase were somehow increased in the dark‐treated ht1‐9 and cbc1 cbc2 leaves (Fig. S8; Dk). Lack of these kinases' functions might have an indirect influence on PM H+‐ATPase phosphorylation, although mis‐regulation of PM anion channels in the mutants is likely to antagonise PM H+‐ATPase to prevent stomatal opening (Hosotani et al., 2021). Mesophyll cells also undergo photosynthesis‐dependent PM H+‐ATPase phosphorylation under light; however, dark‐induced dephosphorylation in mesophyll cells is slower than in guard cells, even though carbonic anhydrases are expressed in both types of cell (Hu et al., 2010; Okumura et al., 2016). This suggests that rapid dephosphorylation in guard cells may be mediated by guard‐cell‐rich components. HT1 and CBCs belong to the family of Raf‐like kinases (Hayashi et al., 2020). Another Raf‐like kinase expressed in guard cells, BHP regulates the blue light‐induced phosphorylation of guard‐cell PM H+‐ATPase in isolated epidermal fragments and GCPs (Hayashi et al., 2017). Interestingly, we found that the phosphorylation level of guard‐cell PM H+‐ATPase in the leaves of a bhp‐1 knock‐out mutant was lower than that of the wild‐type, regardless of the light conditions (Fig. S9). This suggests that BHP could be a negative regulator of rapid guard‐cell PM H+‐ATPase dephosphorylation, which should be investigated in future studies.

Recent biochemical and live‐imaging analyses indicated that CO2 does not trigger an increase in ABA levels or activate OST1 in guard cells; therefore, CO2‐induced stomatal closure may be independent of ABA signalling and stimulated downstream of OST1 (Hsu et al., 2018; Zhang et al., 2020). Given that dark‐induced guard‐cell PM H+‐ATPase dephosphorylation occurs in response to CO2 levels, the occurrence of dark‐induced guard‐cell PM H+‐ATPase dephosphorylation in ost1‐3 suggests that it may not be a downstream target of OST1 during dark‐induced stomatal closure (Fig. S10). A recent study suggested that ABA is not essential in dark‐induced stomatal closure (Pridgeon & Hetherington, 2021). Presence of dark‐induced dephosphorylation of PM H+‐ATPase in ost1‐3 implies that ABA might not be essential in the dephosphorylation. Future studies could be expanded to other ABA signalling components to elucidate this.

Rapid PM H+‐ATPase dephosphorylation did not occur in the guard cells of mutants pp2c.d6 or pp2c.d6/9 following the light–dark transition; however, guard‐cell PM H+‐ATPase of pp2c.d6 was dephosphorylated when maintained in dark conditions (Fig. 5a). Similarly, pp2c.d6/9 exhibited delayed stomatal closure in the dark, while stomatal closure was barely affected in pp2c.d6 (Fig. 5c–e). Taken together, our results suggested that PP2C.D6 initiates dephosphorylation upon light–dark transition and PP2C.D9 then maintains the PM H+‐ATPase in a dephosphorylated state to promote rapid stomatal closure. A recent study indicated that PP2C.D2 and D5, as well as D6, regulate stomatal movement (Wong et al., 2021). The role of PP2C.D isoforms in dark‐induced PM H+‐ATPase dephosphorylation in guard cells should be explored in future studies. In addition, how PP2C.Ds are regulated by dark conditions and/or CO2 is currently unknown. PP2C.Ds are inhibited by SMALL AUXIN UP RNAs (SAURs), which in turn regulate PM H+‐ATPase phosphorylation in seedlings (Spartz et al., 2014). Notably, stomatal opening is suppressed in saur56 saur60 (Wong et al., 2021). The involvement of SAURs in rapid dark‐induced PM H+‐ATPase dephosphorylation in guard cells should be investigated in future studies.

Based on the results of this study, we propose a new model of stomatal closure in leaves involving CO2‐induced rapid PM H+‐ATPase dephosphorylation, which occurs alongside CO2‐mediated activation of PM anion channels in guard cells (Fig. 6). Although the genetic relationship between carbonic anhydrases and PP2C.D6/9 should be confirmed in future work, our findings highlighted how guard‐cell PM H+‐ATPase, a driver of stomatal opening, is regulated in response to the dark or CO2, leading to stomatal closure in leaves. This study also reports a novel mechanism of plant responses to CO2. eCO2 and NaHCO3 treatments induced dephosphorylation even in the light (Figs 3b, S4). Interestingly, NaHCO3‐induced dephosphorylation under blue light was slightly delayed compared with that in the dark (Fig. S4). This delay may be explained by competition between blue light‐induced phosphorylation and CO2‐meidated dephosphorylation of PM H+‐ATPase in guard cells. eCO2‐induced dephosphorylation under RL was also slightly delayed and required higher level of C i compared with the dephosphorylation upon the light–dark transition (Fig. 3b). This may imply that red‐light‐induced phosphorylation of guard‐cell PM H+‐ATPase is a C i‐independent response and competes with the dephosphorylation. This hypothesis would be consistent with the idea that red‐light‐induced stomatal opening is mediated both by C i‐dependent and C i‐independent mechanisms (Matrosova et al., 2015). Therefore, we consider that the phosphorylation level of guard‐cell PM H+‐ATPase may be regulated both by C i ‐dependent dephosphorylation and C i‐independent light‐induced phosphorylation (Fig. 6). Furthermore, the model suggests that fluctuations in photosynthetic activity during the daytime, which occurs due to the fluctuating light conditions (Tanaka et al., 2019), may be a matter for the regulation of stomatal movement in the field. Rapid stomatal closure might prevent deterioration in plant water use efficiency when the photosynthetic capacity is lowered. Future investigations could explore how stomatal movement is regulated in unstable light environments.

Fig. 6.

Fig. 6

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Putative model of the early processes of stomatal closure in leaves, involving CO2 ‐mediated regulation of plasma membrane (PM) H+‐ATPase in guard cells. The increase in intercellular CO2 concentration (C i ↑), which is caused by the cessation of photosynthesis (A ↓) under the light–dark transition or elevated CO2 treatment, induces the dephosphorylation of PM H+‐ATPase, which would prevent 14‐3‐3 binding and lead to inactivation of PM H+‐ATPase (Kinoshita & Shimazaki, 2002). The dephosphorylation probably requires the activity of carbonic anhydrases CA1 and CA4, and may be catalysed at least by PP2C.D6 and PP2C.D9. Blue light (BL)‐ and red light (RL)‐induced phosphorylation of PM H+‐ATPase (Kinoshita & Shimazaki, 1999; Ando & Kinoshita, 2018), which are likely to be independent of C i, may compete against the dephosphorylation. Along with carbonic anhydrase‐mediated activation of PM anion channels, inactivation of PM H+‐ATPase by dephosphorylation may efficiently induce PM depolarization in guard cells, prompting the release of solutes from the cells and subsequent stomatal closure (Roelfsema et al., 2012; Zhang et al., 2018; Dubeaux et al., 2021). Red lines show the signal transduction pathways suggested in this study. Dotted lines indicate omitted or unidentified signalling components. Arrows and T‐bars represent positive and negative regulation, respectively.

Author contributions

EA, HK and TK designed the research; EA, KF and TK performed the research; and EA, HK, KF, TK and IT wrote the article.

Supporting information

Fig. S1 Guard cells in isolated epidermal fragments lack rapid responses to dark conditions.

Fig. S2 Dark‐induced stomatal closure is abolished in Arabidopsis thaliana ost2‐1 leaves.

Fig. S3 Ambient CO2 concentration (C a), photosynthetic rate (A), intercellular CO2 concentration (C i) and stomatal conductance (g s) in Arabidopsis thaliana wild‐type Col‐0 leaves recorded in the experiment shown in Fig. 3(a–c).

Fig. S4 Blue light slightly delays NaHCO3‐induced dephosphorylation of guard‐cell plasma membrane H+‐ATPase in Arabidopsis thaliana wild‐type Col‐0 epidermal fragments.

Fig. S5 NaHCO3‐induced dephosphorylation of plasma membrane H+‐ATPase in Arabidopsis thaliana wild‐type Col‐0 guard‐cell protoplasts.

Fig. S6 Elevated CO2 (eCO2)‐induced dephosphorylation of guard‐cell plasma membrane H+‐ATPase is impaired in Arabidopsis thaliana ca1 ca4 and pp2c.d6/9 leaves.

Fig. S7 Stomatal density of Arabidopsis thaliana leaves used for the gas‐exchange analyses.

Fig. S8 HT1, CBC1 and CBC2 are not likely to function as a negative regulator of dark‐induced dephosphorylation of guard‐cell plasma membrane H+‐ATPase in Arabidopsis thaliana leaves.

Fig. S9 The phosphorylation of guard‐cell plasma membrane H+‐ATPase is suppressed in Arabidopsis thaliana bhp‐1 leaves under all light conditions tested.

Fig. S10 Dark‐induced dephosphorylation of guard‐cell plasma membrane H+‐ATPase is not impaired in Arabidopsis thaliana ost1‐3 leaves.

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.

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Acknowledgements

Seeds of ca1 ca4 were kindly provided by Dr Julian I. Schroeder (University of California, San Diego). Seeds of cbc1 cbc2 and ht1‐9 were kindly provided by Dr Ken‐ichiro Shimazaki (Kyushu University). We thank Dr Masaru Kono (The University of Tokyo) for technical advice. We also thank Ms Kyomi Taki (Nagoya University) for technical assistance. This work was supported by a Grant‐in‐Aid for Japan Society for the Promotion of Science Research Fellow (grant no. 20 J00392 to EA) and Grants‐in‐Aid for Scientific Research from the Ministry of Education, Culture, Sports, Science and Technology, Japan (grant nos. 20H05687 and 20H05910 to TK).

Contributor Information

Eigo Ando, Email: e.ando.ac@gmail.com.

Toshinori Kinoshita, Email: kinoshita@bio.nagoya-u.ac.jp.

Data availability

The data that support the findings of this study are available from TK upon reasonable request.

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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 Guard cells in isolated epidermal fragments lack rapid responses to dark conditions.

Fig. S2 Dark‐induced stomatal closure is abolished in Arabidopsis thaliana ost2‐1 leaves.

Fig. S3 Ambient CO2 concentration (C a), photosynthetic rate (A), intercellular CO2 concentration (C i) and stomatal conductance (g s) in Arabidopsis thaliana wild‐type Col‐0 leaves recorded in the experiment shown in Fig. 3(a–c).

Fig. S4 Blue light slightly delays NaHCO3‐induced dephosphorylation of guard‐cell plasma membrane H+‐ATPase in Arabidopsis thaliana wild‐type Col‐0 epidermal fragments.

Fig. S5 NaHCO3‐induced dephosphorylation of plasma membrane H+‐ATPase in Arabidopsis thaliana wild‐type Col‐0 guard‐cell protoplasts.

Fig. S6 Elevated CO2 (eCO2)‐induced dephosphorylation of guard‐cell plasma membrane H+‐ATPase is impaired in Arabidopsis thaliana ca1 ca4 and pp2c.d6/9 leaves.

Fig. S7 Stomatal density of Arabidopsis thaliana leaves used for the gas‐exchange analyses.

Fig. S8 HT1, CBC1 and CBC2 are not likely to function as a negative regulator of dark‐induced dephosphorylation of guard‐cell plasma membrane H+‐ATPase in Arabidopsis thaliana leaves.

Fig. S9 The phosphorylation of guard‐cell plasma membrane H+‐ATPase is suppressed in Arabidopsis thaliana bhp‐1 leaves under all light conditions tested.

Fig. S10 Dark‐induced dephosphorylation of guard‐cell plasma membrane H+‐ATPase is not impaired in Arabidopsis thaliana ost1‐3 leaves.

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Data Availability Statement

The data that support the findings of this study are available from TK upon reasonable request.

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