Phosphorylation of WDR48 by phototropins drives starch degradation to promote stomatal opening

Domain: chemistry
Published: 2026-03-06
Editorial outcome: minor_revision

Abstract
Plant stomata open in response to blue light under a background of red light. Red light induces starch accumulation through guard cell photosynthesis, whereas blue light causes rapid starch degradation via the activation of plasma membrane H+-ATPase through phosphorylation of BLUS1 by phototropins, thus promoting stomatal opening. However, how phototropins mediate starch degradation remains unclear. In this study, we identified WD-repeat protein 48 (WDR48), which is essential for starch degradation. Phosphoproteomic analysis revealed that WDR48 was phosphorylated in response to blue light. Mutation of WDR48 impaired starch degradation and stomatal opening by blue light. Phototropins interacted with and phosphorylated WDR48. WDR48 and BLUS1 constituted separate signalling pathways required for starch degradation. We propose that the coordinated control of starch synthesis by red light and its degradation by blue light signalling through WDR48 and BLUS1 is a key mechanism for stomatal opening by red and blue light.

Introduction
Plant stomata, which are formed by a pair of guard cells, are critical in regulating gas exchange between plants and the atmosphere. These specialised cells adjust stomatal pore size in response to changing environmental conditions1–3. Light triggers stomatal opening, facilitating the uptake of carbon dioxide (CO ) for photosynthesis as well as the transport of water and nutrients from the roots via transpiration. Stomatal responses to light can be divided into photosynthesis-dependent and blue light-dependent responses2,4. The photosynthesis-dependent response, which can be activated by red light, is induced by continuous high fluence rates and eliminated by 3-(3,4-dichlorophenyl)-1,1-dimethylurea (DCMU), a photosynthetic electron transport inhibitor5,6. Guard cells and mesophyll cells contain photosynthetically active chloroplasts, contributing to red light-induced stomatal opening7–9. In contrast, the blue light response occurs at low fluence rates and relies on photoreceptor phototropin-coupled ion transport and metabolic rearrangements in the guard cells10,11.

Furthermore, an interaction occurs between photosynthesis- and blue light-dependent responses. The blue light response is enhanced when blue light is superimposed on a strong red light background and simultaneous irradiation with blue and red light results in greater stomatal opening than irradiation with blue or red light alone2,12–16. The synergistic effect of blue and red light on stomatal opening may be important for guard cells to integrate information from blue light and photosynthesis to appropriately control stomatal opening and CO uptake under photosynthetically active conditions. Studies have suggested that both blue light signals and red light-driven mesophyll cell photosynthesis activate the plasma membrane H⁺-ATPase and inactivate S-type anion channels, thereby contributing synergistically to stomatal opening in intact leaves17-19. This synergistic effect of blue and red light on stomatal opening has also been observed in isolated epidermis, suggesting the presence of mesophyll cell-independent regulation15. However, the underlying mechanisms remain elusive.

Phototropins are plant-specific blue light receptor kinases containing two light, oxygen, or voltage domains (LOV1 and LOV2) at the N-terminus, each of which binds a flavin mononucleotide (FMN) as a chromophore20. Light perception, mainly through the LOV2 domain, activates the C-terminal kinase domain and induces receptor autophosphorylation21. Light-activated phototropins phosphorylate their substrate BLUE LIGHT SIGNALING1 (BLUS1), a guard cell-specific Ser/Thr protein kinase22,23. BLUS1 contains a C-terminal regulatory domain that represses the N-terminal kinase domain. Phototropins phosphorylate a conserved Ser residue within the regulatory domain of BLUS1, alleviating suppression of the kinase domain16. The activated BLUS1 transduces signals via downstream elements such as a Raf-like MAP kinase kinase kinase (MAPKKK) BLUE LIGHT-DEPENDENT H+-ATPASE PHOSPHORYLATION (BHP) and type 1 protein phosphatase (PP1), ultimately activating the plasma membrane H+-ATPase via the phosphorylation of two Thr residues within the C-terminal autoinhibitory domain17,18,24-26. In addition to BLUS1, phototropins phosphorylate a Raf-like MAPKKK CONVERGENCE OF BLUE LIGHT AND CO 1 (CBC1)27. CBC1 and its paralogue CBC2 inhibit plasma membrane S-type anion channels in response to blue light27,28. Consequent H+-ATPase activation and anion channel inhibition drive plasma membrane hyperpolarisation, providing the driving force for K+ influx through voltage-gated inward K+ channels29. K+ accumulation drives water uptake, causing swelling of guard cells and stomatal opening.

Simultaneously with the regulation of membrane potential, phototropins induce rapid starch degradation in guard cell chloroplasts30. Chloroplasts are a predominant feature of guard cells as they are not found in other leaf epidermal cells. Guard cells partially synthesise starch via autonomous photosynthetic activity31,32. The degradation products of starch by blue light are converted to glucose, contributing to maintaining cytosolic sugar homeostasis needed for stomatal opening30,33. Starch degradation products have also been suggested to serve as a precursor of malate, which acts as a counter-ion to K+ influx2,34. Mutations of BLUS1 or AUTOINHIBITED H+-ATPASE 1 (AHA1), a major isoform of plasma membrane H+-ATPase in guard cells, impairs starch degradation by blue light, revealing that BLUS1-mediated H+-ATPase activation is involved in this process30,35. Similarly, mutation of the starch degradation enzymes -AMYLASE 1 (BAM1) and -AMYLASE 3 (AMY3) impairs blue light-induced starch degradation and stomatal opening30. However, the mechanisms by which phototropins cause rapid starch degradation in guard cell chloroplasts remain unclear.

In this study, we performed phosphoproteomic analysis to uncover novel factors involved in phototropin-controlled stomatal opening. We identified a novel phototropin kinase substrate, WD-repeat protein 48 (WDR48), that is essential for starch degradation and stomatal opening by blue light. WDR48 and BLUS1 form separate signalling pathways implicated in blue light-dependent starch degradation in guard cells.

Results
WDR48 is phosphorylated in response to blue light and regulates stomatal opening
We conducted a phosphoproteomic analysis using Arabidopsis thaliana guard cell protoplasts from the wild type and the phot1-5 phot2-1 mutant, and analysed proteins whose phosphorylation levels changed in response to blue light in the wild type but not in the phot1-5 phot2-1 mutant17,22,27. We identified a WD-repeat protein (WDR), At3g05090, known as LATERAL ROOT STIMULATOR 1 (LRS1)36, which was phosphorylated in a blue light- and phototropin-dependent manner (Fig. 1a, b). This WDR protein is conserved amongst eukaryotes and is generally referred to as WDR4837. We, therefore, refer to this protein as WDR48 in this study. WDR48 contains six WD40 repeats at the N-terminus and a RAWUL domain at the C-terminus that appears to function in protein–protein interactions37. Ser-393, which is located between the two domains, was identified as a blue light-dependent phosphorylation site (Fig. 1c).

We generated phospho-specific antibodies against Ser-393 of WDR48 and found that Ser-393 was weakly phosphorylated under red light, whereas phosphorylation was enhanced upon blue light irradiation in the wild type and absent in the phot1-5 phot2-1 mutant (Fig. 1d, e). The bands observed under both red and blue light conditions disappeared after λ-protein phosphatase (λ-PPase) treatment in the wild type (Supplementary Fig. 1a, b) and were absent in both the wdr48-1 mutant and the phosphodefective S393A mutant described later (Supplementary Fig. 1c–f), suggesting that the phospho-specific antibodies generated in this study specifically recognise phosphorylated WDR48.

To determine the physiological role of WDR48 in the stomatal opening, we obtained two T-DNA insertion mutants, wdr48-1 (GABI_585_E01) and wdr48-2 (SALK_059570C), harbouring a T-DNA insertion in the 2nd and 14th introns, respectively (Fig. 1f). Immunoblot analysis using anti-WDR48 antibodies revealed that both wdr48 mutants were null alleles (Fig. 1g). We confirmed that there were no significant differences in stomatal size or density between the wild-type and wdr48 mutants (Supplementary Fig. 2). We then examined the stomatal opening in the epidermal peels, and wdr48-1 and wdr48-2 plants exhibited defects in the stomatal opening in response to blue light, similar to the phot1-5 phot2-1 mutant (Fig. 1h).

To examine whether the wdr48 mutant impairs the activation of plasma membrane H+-ATPase by blue light, we measured H+ pumping in guard cell protoplasts. The wdr48-1 mutant exhibited the same level of blue light-dependent H+ pumping activity as the wild-type (Fig. 1i, j). The phosphorylation of H+-ATPase in response to blue light was not affected in the wdr48-1 mutant (Fig. 1k, l). The electrophoretic mobility shift on SDS-PAGE confirmed the blue light-dependent autophosphorylation of phototropins in the wdr48-1 mutant (Fig. 1m). Furthermore, blue light-dependent phosphorylation of BLUS1 was prominent in the wdr48-1 mutant (Fig. 1n). Overall, these results suggest that WDR48 stimulates stomatal opening by regulating mechanisms that are distinct from H+-ATPase activation.

WDR48 regulates stomatal opening via the regulation of starch dynamics in guard cells
To determine whether wdr48 impairs blue light-dependent starch degradation, we quantified the starch in guard cells using the pseudo-Schiff propidium iodide (PS-PI) staining technique30,38. We prepared leaf epidermis from dark-adapted wild-type plants and observed little starch accumulation in guard cells in the dark (Fig. 2a, b). Irradiation of the epidermis with red light induced starch accumulation, and subsequent exposure to blue light caused starch degradation. The application of DCMU to the epidermis suppressed starch accumulation by red light and starch degradation by blue light (Fig. 2a, b). The wdr48-1 mutant showed starch accumulation in guard cells under red light but failed to cause starch degradation by blue light, similar to the phot1-5 phot2-1 mutant (Fig. 2c, d).

We determined the time course of starch degradation and stomatal opening in response to blue light. When wild-type epidermis was irradiated with blue light following red light treatment to ensure starch accumulation, guard cell starch was rapidly degraded within 5 min (Fig. 2e, f). Consistent with starch degradation, the stomatal opening was prominent 5 min after blue light irradiation and peaked at approximately 10 min (Fig. 2g). In contrast, no starch degradation or stomatal opening was detected in phot1-5 phot2-1 and wdr48-1 mutants after 30 min of blue light irradiation (Fig. 2e–g). These findings suggest that WDR48 controls phototropin-mediated starch degradation to allow rapid stomatal opening in response to blue light.

Phototropins also regulate chloroplast movements in addition to stomatal opening20. Because WDR48 controls starch degradation in chloroplasts, we examined whether chloroplast movements are impaired in the wdr48 mutants using the green and white band assays39. The wdr48 mutants exhibited chloroplast accumulation under weak blue light and avoidance under strong blue light, similar to the wild type (Supplementary Fig. 3), suggesting that WDR48 is unlikely to function in chloroplast movements.

Phosphorylation of WDR48 is necessary for starch degradation and stomatal opening
To examine the physiological role of blue light-dependent phosphorylation of WDR48 in stomatal responses, we generated transgenic plants expressing phosphodefective or phosphomimetic forms of WDR48, in which Ser-393 was replaced with Ala (S393A) or Glu (S393E), respectively, as an N-terminal fusion to GFP in the wdr48-1 mutant background under the control of its promoter. We selected transgenic lines that showed comparable levels of expression to endogenous WDR48 by immunoblot analysis using anti-WDR48 antibodies (Supplementary Fig. 4). Expression of GFP-WDR48 and GFP-S393E complemented the inhibition of starch degradation and stomatal opening in response to blue light in the wdr48-1 mutant (Fig. 3a–c). In contrast, when GFP-S393A was introduced into the wdr48-1 mutant, neither starch degradation nor stomatal opening was restored.

We evaluated the stomatal opening in intact leaves using a gas exchange system. Strong red light irradiation increased stomatal conductance as a result of photosynthesis-dependent stomatal opening, and the superimposition of weak blue light further promoted stomatal opening (Fig. 3d). When compared to the wild-type plants, both red light- and blue light-dependent stomatal opening were diminished in the wdr48-1 mutant (Fig. 3e, f). Furthermore, GFP-WDR48 and GFP-S393E lines exhibited enhanced stomatal conductance in response to red and blue light. In contrast, the phosphodefective GFP-S393A line complemented the wdr48-1 stomatal phenotype under red light but failed to restore blue light-dependent stomatal opening. These findings suggest that phosphorylation of WDR48 is crucial for starch degradation and stomatal opening in response to blue light, and that WDR48 also plays a role in regulating stomatal opening under red light via a phosphorylation-independent mechanism.

In addition to its role in stomatal opening, WDR48 has been shown to be implicated in lateral root formation36. In the wdr48 mutants, both lateral root formation and primary root growth were impaired (Supplementary Fig. 5). These defects were rescued not only by expression of wild-type WDR48 but also by phosphodefective and phosphomimetic forms of WDR48, suggesting that WDR48 regulates root development through a function that is independent of its phosphorylation.

WDR48 is a direct substrate of phototropin kinases
We further explored the mechanism of WDR48 phosphorylation by phototropins. Blue light-dependent phosphorylation of WDR48 was detected in guard cell protoplasts in the wild type but not in the phot1-5 phot2-1 double mutant (Fig. 4a, b). WDR48 phosphorylation was also detected in the phot1-5 and phot2-1 single mutants, indicating that phot1 and phot2 mediate WDR48 phosphorylation. Consistent with this finding, starch granules were degraded in the phot1-5 and phot2-1 mutants but not in the phot1-5 phot2-1 mutant (Supplementary Fig. 6a, b). We evaluated the phosphorylation of WDR48 in mutants of BLUS1 and CBC kinases, which are phosphorylation substrates of phototropins22,27. WDR48 phosphorylation by blue light was prominent in the blus1-1 and cbc1-1 cbc2-1 mutants (Fig. 4c, d), indicating that WDR48 phosphorylation is mediated by blue light signalling separate from the BLUS1 and CBC kinases.

We then analysed the phosphorylation time course of WDR48 in greater detail. WDR48 was phosphorylated 15 s after blue light pulse irradiation, peaked at 2 min, and was dephosphorylated within 20 min (Fig. 4e, f). The phosphorylation time course of WDR48 was consistent with that of BLUS1 (Fig. 4e, f). Therefore, we hypothesised that WDR48 is a phosphorylation substrate of phototropins, such as BLUS1 and CBC1. To test this possibility, we produced recombinant FLAG- WDR48 and His-phot1 or His-phot2 proteins using an in vitro transcription/translation system and performed an in vitro kinase assay. Phosphorylation of WDR48 at Ser-393 was detected when FLAG-WDR48 was incubated with His-phot1 or His-phot2 under blue light, but not in darkness (Fig. 4g, h, Supplementary Fig. 7a, b). Blue light-dependent phosphorylation of FLAG-WDR48 was not observed when incubated with the kinase-dead forms His-phot1 (D806N) or His-phot2 (D720N). Furthermore, λ-PPase treatment abolished the blue light-dependent phosphorylation of FLAG-WDR48 by His-phot1 or His-phot2 (Supplementary Fig. 7c–f).

The in vivo interaction between WDR48 and phot1 was confirmed by a co-immunoprecipitation assay using transgenic plants expressing 3×FLAG-WDR48. Endogenous phot1 protein was co-precipitated with 3×FLAG-WDR48 irrespective of blue light irradiation (Fig. 4i, j). We further verified direct binding between FLAG-WDR48 and HA-phot1 by an in vitro pull-down assay, independent of blue light (Fig. 4k, l). To identify the region of phot1 responsible for the interaction, we performed an in vitro pull-down assay with FLAG-phot1 fragments. His-WDR48 bound to the C-terminal kinase domain of phot1 (FLAG-P1C) but not to the N-terminal LOV1 and LOV2 domains (FLAG-P1N) (Fig. 4m), indicating that the kinase domain mediates the interaction with WDR48. From these results, we concluded that WDR48 is a direct target of phototropin kinases.

WDR48 interacts with phot1 at the plasma membrane and chloroplast envelope
The subcellular localisation of phot1-GFP and GFP-WDR48 in guard cells was observed using confocal laser scanning microscopy. In the dark, GFP fluorescence of phot1-GFP was mainly observed at the plasma membrane, with weaker signals occasionally detectable around chloroplasts; meanwhile, blue light irradiation enhanced phot1-GFP signals in the cytoplasm (Fig. 5a, b). In contrast, the GFP-WDR48 signals were partially aggregated and distributed in the cytoplasm, with additional signals detected along the plasma membrane and the chloroplast envelope, and blue light irradiation did not significantly alter its localisation patterns (Fig. 5a, b, Supplementary Fig. 8). Furthermore, neither the phosphodefective nor the phosphomimetic mutations altered the localisation of GFP-WDR48 (Supplementary Fig. 9). To elucidate the localisation of phot1-GFP and GFP-WDR48 on the chloroplast envelope more precisely, we introduced the chloroplast envelope marker OUTER ENVELOPE MEMBRANE PROTEIN 7 (OEP7)-mCherry into plants expressing either phot1-GFP or GFP-WDR4840. Analysis revealed that signals from both proteins were detectable on the chloroplast envelope under both dark and blue light conditions, as confirmed by colocalisation with OEP7-mCherry (Supplementary Fig. 10).

We further visualised the interaction between phot1 and WDR48 in plants using a bimolecular fluorescence complementation (BiFC) assay. When WDR48 fused to the N-terminal half of YFP (YFPn-WDR48) was co-expressed with phot1 fused to the C-terminal half of YFP (YFPc-phot1) in mesophyll cell protoplasts, YFP signals were detected at the cell periphery and co-localised with the signals of POTASSIUM CHANNEL IN ARABIDOPSIS THALIANA 1 (KAT1)-mCherry, a plasma membrane marker (Fig. 5c)41. In addition, when the depth of focus was changed, YFP signals were detected around the chloroplasts, where they co-localised with OEP7-mCherry (Fig. 5e). Furthermore, to confirm the interaction between phot1 and WDR48 in the BiFC assay, we performed a BiFC competition assay in which non-fused interacting proteins inhibited the BiFC reaction42. A decrease in the YFP signal was detected when 3×FLAG-WDR48 was co-expressed as a competitor for the BiFC reaction (Fig. 5c–f). We also confirmed that no YFP signals were observed when PIN-FORMED1 (PIN1) fused to the C-terminal half of YFP (YFPc-PIN1) was co-expressed with YFPn-WDR48 as a negative control (Fig. 5c). These findings suggest that WDR48 and phot1 interact at the plasma membrane and chloroplast envelope.

BLUS1 and WDR48 coordinately regulate starch degradation in response to blue light
Blue light signalling mediated by BLUS1 activates the plasma membrane H+-ATPase, which is required for starch degradation during stomatal opening (Supplementary Fig. 6c, d)30. Considering that the blue light activation of H+-ATPase was not affected in the wdr48 mutant, WDR48 is most likely not involved in signalling upstream of H+-ATPase activation. Furthermore, phosphorylation of WDR48 was induced by blue light but not by the application of fusicoccin (Fc), an activator of H+-ATPase (Supplementary Fig. 11a, b). In addition, blue light-dependent WDR48 phosphorylation was observed in the blus1-1 and aha1-9 mutants, as in the wild type (Fig. 4c, d; Supplementary Fig. 11c, d), suggesting that WDR48 does not function downstream of H+-ATPase activation. Thus, WDR48 appears to constitute a separate signalling pathway from that of H+-ATPase activation and induce starch degradation.

To further confirm that WDR48 induces starch degradation via a pathway distinct from the activation of H+-ATPase, we examined starch degradation by Fc. A previous study indicated that treatment with 10 µM Fc induced starch degradation in guard cells in the isolated epidermis, regardless of blue light irradiation30. Consistent with this finding, treatment with 10 µM Fc under red light induced starch degradation and stomatal opening in the wild-type as well as in the wdr48 mutants (Supplementary Fig. 12a–c). However, treatment with such a high concentration (10 µM) of Fc caused a two-times higher rate of H+ pumping compared to blue light conditions (Supplementary Fig. 12d) and induced stomatal opening even in the dark without starch degradation (Supplementary Fig. 12e–g). However, when epidermal peels were treated with 1 µM Fc, which induced a similar rate of H+ pumping to that of blue light (Supplementary Fig. 12d), neither starch degradation nor stomatal opening was observed under red light or in the dark (Supplementary Figs. 12a–c and 12e–g). These results suggest that activation of H+-ATPase alone is not sufficient to trigger starch degradation and stomatal opening under physiological conditions and that additional blue light signals are required for the induction of these responses.

Next, we investigated starch degradation and stomatal opening by blue light and 1 µM Fc in blus1 and wdr48 single mutants and blus1 wdr48 double mutant. If BLUS1 and WDR48 function in separate signalling pathways and both pathways are required for starch degradation, we would expect that the application of 1 µM Fc, together with blue light, induce starch degradation and stomatal opening in the blus1 mutant, but not in the wdr48 or blus1 wdr48 mutants. In the wild-type, blue light irradiation or simultaneous application of blue light and Fc induced starch degradation and stomatal opening, but Fc alone did not induce either response (Fig. 6a–c). In contrast, blue light and Fc did not stimulate starch degradation or stomatal opening in the wdr48 mutant under any condition. Similarly, in the blus1 mutant, which lacks blue light-dependent activation of H+-ATPase22, neither blue light nor Fc alone induced starch degradation or stomatal opening, whereas the combination of blue light and Fc elicited both responses. However, blue light- and Fc-induced starch degradation and stomatal opening observed in the blus1 mutant were not found in the blus1 wdr48 double mutant.

Assuming that both H+-ATPase activation and WDR48 phosphorylation are required for starch degradation, the phosphodefective WDR48 variant would not exhibit starch degradation even when H+-ATPase is activated by Fc, because WDR48 cannot be phosphorylated. In contrast, the phosphomimetic WDR48 variant is expected to trigger starch degradation if H+-ATPase is activated by Fc. To test this, we examined starch degradation and stomatal opening in transgenic lines expressing either phosphodefective (GFP-S393A) or phosphomimetic (GFP-S393E) variants of WDR48, alongside the GFP-WDR48 line and the wdr48 mutant. As expected, starch degradation and stomatal opening were induced by blue light or by the combination of blue light and Fc in the GFP-WDR48 line (Fig. 6d–f). In contrast, neither response was observed under any treatment in the phosphodefective GFP-S393A line or in the wdr48 mutant. Notably, in the phosphomimetic GFP-S393E line, Fc treatment alone was sufficient to induce both starch degradation and stomatal opening in the absence of blue light.

In transgenic lines expressing the phosphomimetic GFP-S393E in phot1 phot2 wdr48 or blus1 wdr48 mutant backgrounds, blue light alone did not induce starch degradation or stomatal opening, which is consistent with impaired H⁺-ATPase activation in these mutants (Fig. 6g–i). However, when Fc was applied, both responses were successfully triggered. In the GFP-S393E expressing line in the aha1 wdr48 background, blue light alone did not induce starch degradation or stomatal opening, and neither Fc nor blue light in combination with Fc induced these responses. These findings suggest that phosphorylation of WDR48, together with BLUS1-mediated H+-ATPase activation is essential for starch degradation and stomatal opening.

Methods
Plant materials and growth conditions
Arabidopsis thaliana accession Columbia-0 (Col-0) was used as the wild-type. phot1-5 phot2-110, phot1-551, phot2-152, blus1-122, aha1-935, and cbc1-1 cbc2-127 have been described previously. T-DNA insertion mutants of wdr48-1 (GABI_585_E01) and wdr48-2 (SALK_059570C) were obtained from the Nottingham Arabidopsis Stock Centre (NASC). The transgenic plants of phot1-GFP have been described previously22. Plants were grown on a soil:vermiculite mixture (1:1) for 4 weeks under white light (50 µmol m-2 s-1) with 14 h/10 h light/dark cycles at 24 ℃.

Phosphoproteomic analysis
Phosphoproteomic analysis was conducted using guard cell protoplasts from the wild-type and phot1-5 phot2-1 mutants17,22,27. Guard cell protoplasts were incubated under red light (600 µmol m-2 s-1) for 30 min, after which a pulse of blue light (100 µmol m-2 s-1, 30 s) was superimposed on the background of red light. The reaction was terminated 2.5 min after the start of blue light illumination by the addition of trichloroacetic acid to the protoplast suspension. Following digestion with Lys-C and trypsin, the phosphopeptides were enriched by hydroxy acid-modified metal oxide chromatography (HAMMOC) using lactic acid-modified titania53,54. The enriched phosphopeptides were analysed by nanoLC/MS/MS in Data Dependent Acquisition (DDA) mode using LTQ-Orbitrap (Thermo Fisher Scientific)17,22,27 or Orbitrap Fusion Lumos (Thermo Fisher Scientific) under the following conditions.

The Orbitrap Fusion Lumos was coupled with Ultimate3000 pump (Thermo Fisher Scientific) and an HTC-PAL autosampler (CTC Analytics). Reprosil-Pur 120 C18-AQ beads (1.9 μm; Dr. Maisch GmbH) packed in a self-pulled needle (150 mm length × 100 μm i.d., 6-μm opening) were used as the nanoLC column. The mobile phases consisted of 0.5% acetic acid (A) and 0.5% acetic acid and 80% acetonitrile (B). A three-step linear gradient of 5% to 10% B in 5 min, 10% to 40% B in 60 min, 40% to 100% B in 5 min, and 100% B for 10 min was applied at a flow rate of 500 nL/min. A spray voltage of 2,400 V was applied, and the FAIMS compensation voltage was set to -40, -60, and -80. The MS scan range was m/z 300–1500. MS scans were performed using the Orbitrap with r = 120,000 and subsequent MS/MS scans were performed by the quadrupole ion trap. Synchronous precursor selection (SPS) was employed with a higher-energy collisional dissociation (HCD) at 10 SPS precursors. The HCD collision energy was set to 30.

The raw LTQ-Orbitrap data were analysed with Mascot version 2.3 (Matrix Science) against the TAIR database (release 10) as previously described17,22,27. The raw Orbitrap Fusion Lumos data were analysed with MSFragger (version 4.1)55 within the FragPipe platform (v22.0) against the TAIR database (release 10) as follows. Mass tolerances of 10 ppm and 0.4 Da were applied to precursor and fragment ions, respectively, and mass calibration and parameter optimisation were enabled. Trypsin was specified as the reference enzyme, and up to two missed cleavages were allowed. Carbamidomethylation of cysteine was set as a fixed modification, and oxidation of methionine and phosphorylation of serine, threonine and tyrosine were allowed as variable modifications. PeptideProphet and ProteinProphet in Philosopher56 were used to filter the results with FDR < 1%. The identified phosphopeptides were requantified using Slyline57.

Isolation of guard cell protoplasts and measurement of H+ pumping
Guard cell protoplasts were enzymatically isolated from the leaves25,58. The leaves of 4-week-old plants were homogenised using a blender (Warning), and epidermal fragments were collected on a 58 µm nylon mesh. The epidermal fragments were incubated in an enzyme solution containing 10 mM MES-KOH (pH 5.4), 0.25 M mannitol, 0.5% (w/v) cellulase R-10, 0.05% (w/v) macerozyme R-10, 0.1% (w/v) polyvinylpyrrolidone K-30, 0.2% (w/v) bovine serum albumin, and 1 mM CaCl for 30 min at 24 ℃. The fragments were then recovered on the nylon mesh and washed with 0.3 M mannitol and 1 mM CaCl . Subsequently, the epidermal fragments were incubated in a second enzyme solution containing 10 mM MES-KOH (pH 5.4), 0.4 M mannitol, 1.5% (w/v) cellulase RS, 0.5% (w/v) macerozyme R-10, 0.2% (w/v) bovine serum albumin, and 1 mM CaCl for 1 h at 27 ℃. The guard cell protoplasts were filtered sequentially through 94 µm, 25 µm, and four layers of 10 µm nylon meshes and collected by centrifugation at 420 × g for 18 min. The protoplasts were washed twice with 0.4 M mannitol and 1 mM CaCl and used for analysis. To measure H+ pumping, guard cell protoplasts were incubated in 0.125 mM MES-NaOH (pH 6.0), 1 mM CaCl , 0.4 M mannitol, and 10 mM KCl under red light (300 µmol m-2 s-1) for 2 h, after which a pulse of blue light (100 µmol m-2 s-1) was superimposed on the background of red light for 30 s. H+ pumping was measured using a pH electrode.

Immunoblot analysis
Guard cell protoplasts were incubated in 0.125 mM MES-NaOH (pH 6.0), 1 mM CaCl , 0.4 M mannitol, and 10 mM KCl under red light (300 µmol m-2 s-1) for 30 min, and then a pulse of blue light (100 µmol m-2 s-1) was superimposed on the background of red light for 30 s. The reaction was terminated at 2 min for phot1, phot2, WDR48, and BLUS1 and 3 min for H+-ATPase after the start of irradiation with a blue light pulse by adding trichloroacetic acid to guard cell protoplasts. After washing twice with 50 mM Tris-HCl (pH 8.0), the precipitated proteins were resuspended in the SDS sample buffer and subjected to SDS-SPAGE.

For λ-PPase treatment, guard cell protoplasts were incubated as described above and collected by centrifugation at 10,000 × g for 15 s. Pellets were resuspended in buffer containing 50 mM HEPES (pH 7.5), 100 mM NaCl, 2 mM dithiothreitol (DTT), 0.01% (w/v) Brij 35, 1 mM MnCl , 0.4% (v/v) Triton X-100, 0.5 mM phenylmethylsulfonyl fluoride (PMSF), 10 µM leupeptin, and 400 units of λ-PPase (New England Biolabs, P0753), and incubated at 30 °C for 60 min. After the reaction, SDS sample buffer was added to the λ-PPase-treated guard cell proteins, and the samples were subjected to SDS-PAGE.

Immunoblot analysis was performed as previously reported25,59. Antibodies against WDR48 and phosphorylated Ser-393 of WDR48 were produced by immunising rabbits with recombinant GST-WDR48 (V300-R753) and synthetic phospho-peptide NRARVpSLEGLNPA, respectively. Antibodies against phot1, phot2, BLUS1, pSer348 of BLUST1, H+-ATPase, and pThr948 of AHA1 have been previously described17,22,60,61. Anti-GFP and anti-FLAG-HRP antibodies were purchased from Invitrogen (A-6455) and Sigma-Aldrich (A8592), respectively. Antibodies were used at the following dilutions: 1:1,000 for anti-WDR48, anti-pSer393 of WDR48, and anti-pSer348 of BLUS1; 1:3,000 for anti-BLUS1, anti-GFP, anti-phot1, and anti-phot2; 1:5,000 for anti-pThr948 of AHA1 and anti-FLAG-HRP; and 1:8,000 for anti-H⁺-ATPase.

Measurement of stomatal opening
The stomatal opening in the isolated epidermis was measured as described previously10, with slight modifications. Epidermal strips from leaves of dark-adapted plants were incubated in stomatal opening buffer containing 5 mM MES-bistrispropane (pH 6.5), 50 mM KCl, and 0.1 mM CaCl under red light (50 µmol m-2 s-1) and blue light (10 µmol m-2 s-1) for 2 h at 24 ℃. For the measurements of rapid stomatal opening, the epidermal strips were preirradiated with red light (50 µmol m-2 s-1) for 2 h, after which blue light (10 µmol m-2 s-1) was superimposed on the background of red light or Fc (1 or 10 µM) was added to the buffer, and further incubated for the indicated time. Images of stomata in the abaxial epidermis were acquired using an inverted microscope (Eclipse TS100; Nikon). The stomatal aperture was determined using the ImageJ 1.53q software (National Institutes of Health).

Stomatal conductance in intact leaves was measured using a gas-exchange measurement system (LI-6400; Li-COR), as described previously, with settings of 350 ppm CO , 40–60% relative humidity, 24 °C leaf temperature, and 200 µmol m-1 flow rate62. Leaves of dark-adapted plants were irradiated with red light (300 µmol m-2 s-1) for 1 h, after which blue light (10 µmol m-2 s-1) was superimposed on the background red light for 20 min.

PS-PI stain and quantification of guard cell starch
Abaxial epidermal peels from dark-adapted plants were incubated as described for the measurement of stomatal opening. Guard cell starch granules were fixed and stained with PS-PI staining30,38. Fluorescence images were obtained using a confocal laser scanning microscope (488 nm excitation and 590 nm long pass emission) (Digital Eclipse C1; Nikon).

To quantify the guard cell starch area, we used the X-net63 for automatic starch segmentation. X-net is a cell image segmentation method consisting of two encoders and two decoders and performed better than U-net64. In X-net, the input image is fed into two encoders, and two feature maps in two encoders are aggregated by concatenation. This allows us to extract rich features that cannot be extracted by a single network and improves the segmentation accuracy. To train and evaluate the X-net for starch segmentation, we used 319 images consisting of 2 classes: starch and others. The 319 images were annotated manually using the ImageJ 1.53q software. We cropped the regions of 512 × 512 pixels from the original images to train the X-net because of the memory of the GPU. We used Intersection over Union (IoU), which is the overlap ratio between the segmentation result and manual annotation, as an evaluation measure. X-net was evaluated using four-fold cross-validation. The IoUs of the starch and other classes were 69.75% and 99.63%, respectively. The starch granule area was determined by dividing the labelled pixel area by the pixel density (pixels µm-2) of the analysed image.

Construction of transgenic plants
The 7,370 bp genomic sequence of WDR48 containing the promoter, coding, and 3′-UTR region was amplified using primers 5′-GCTTGATATCGAATTCCGCTTAAGGAAACTAGTGAGTG-3′ and 5′-TAGAACTAGTGGATCCGAGAAGCTAAGTAAACACTAGGC-3′. The resulting product was subcloned into BamH EcoR  site of the pBluescript SK (+) vector (Stratagene) using the In-Fusion HD Cloning Kit (Clontech). Site-directed mutagenesis was performed using a QuikChange Site-Directed Mutagenesis Kit (Stratagene). The primers used were 5′-CTTTTAATAGGGCGAGGGTAGCTTTGGAAGGACTAAATCC-3′ and 5′-GGATTTAGTCCTTCCAAAGCTACCCTCGCCCTATTAAAAG-3′ for WDR48-S393A and 5′-CTTTTAATAGGGCGAGGGTAGAATTGGAAGGACTAAATCC-3′ and 5′-GGATTTAGTCCTTCCAATTCTACCCTCGCCCTATTAAAAG-3′ for WDR48-S393E. To construct the pWDR48:GFP-WDR48 vector, the kanamycin resistance gene, NPT  in pRI 101-AN (TaKaRa), was replaced with the hygromycin resistance gene, HPT, and the resulting vector was designated pPCS201. The GFP sequence was then inserted into the Sal /BamH  sites to generate pPCS202. The upstream 1,398 bp sequence, including the promoter region of WDR48, was amplified using primers 5′-GGCCAGTGCCAAGCTTGCTTAAGGAAACTAGTGAGTG-3′ and 5′-TACCCCCGGGGTCGACTTCAAATTATTCTGAAAGCCTGAAAGGAG-3′, and the product was subcloned into the Hind /Sal  sites of pPCS202. After amplifying the downstream 5,981 bp sequence, including the coding and 3′-UTR regions of WDR48 using primers 5′-CAAGGGATCCGAATTCATGCACCGGGTTGGAAGTGC-3′ and 5′-TGTTGATTCAGAATTCGAGAAGCTAAGTAAACACTAGGC-3′, the resulting product was inserted into the EcoR  site of pPCS202 containing the upstream sequence. To construct the p35S:3×FLAG-WDR48 vector, the 3×FLAG sequence was inserted into the Sal /BamH  sites of pPCS201, and the resulting vector was designated pPCS206. The 5,981 bp genomic sequence, including the coding and 3′-UTR regions of WDR48, was amplified using primers 5′-CGATGACAAGGGATCCATGCACCGGGTTGGAAGTGC-3′ and 5′-TTCAGAATTCGGATCCGAGAAGCTAAGTAAACACTAGGC-3′, and the resulting product was subcloned into BamH  site of pPCS206. Each construct was transformed into the wdr48-1 mutant via floral dip using Agrobacterium tumefaciens strain GV3101.

To construct the pGC1:OEP7-mCherry vector, the NOS promoter and NPT II in pRI 101-AN were replaced with pOLE1:OLE1-TagRFP 65, generating pPCS501. The 35S promoter of pPCS501 was then replaced with the GC1 promoter to yield pPCTS601. OEP7 fused to mCherry was amplified using primers 5′-AAGAAATATGGTCGACATGGGAAAAACTTCGGGAGCG-3′ and 5′-TTCAGAATTCGGATCCTTACTTGTACAGCTCGTCCATG-3′, and the product was subcloned into the Sal /BamH  sites of pPCS601. The construct was introduced into phot1-GFP- or GFP-WDR48-expressing plants.

In vitro pull-down assay
FLAG-WDR48, His-WDR48, FLAG-P1N (N-terminal fragment of phot1, M1-N619), and FLAG-P1C (C-terminal fragment of phot1, M620-F996) were synthesised using an in vitro transcription/translation kit66 (BioSieg). To express HA-phot1, cDNA sequence of PHOT1 fused to an HA tag was amplified using primers 5′-TATAGAATACAAGCTTATGTATCCTTACGATGTTCCAGATTATGCTATGGAACCAACAGAAAAACCATCGAC-3′ and 5′-GCTCGCCCGGGGATCCTCAAAAAACATTTGTTTGCAGATCTTCTAGCTC-3′, and the product was subcloned into the Hind III/BamH I sites of pSP64 poly(A) vector (Promega, P1241). The resulting construct was added to the reaction mixture of the TnT SP6 High-Yield Wheat Germ Protein Expression System (Promega, L3260), supplemented with 10 µM flavin mononucleotide (FMN), as described previously67.

For the in vitro pull-down assay using full-length phot1, FLAG-WDR48 was diluted in buffer containing 20 mM Tris-HCl (pH 7.4), 140 mM NaCl, and 0.1% (v/v) Trinton X-100, and incubated with anti-FLAG M2 affinity gel (Sigma-Aldrich, A2220) at 4 ℃ for 1 h. After washing the beads three times, HA-phot1, diluted in buffer containing 37.5 mM Tris-HCl (pH 7.5), 5.3 mM MgSO , 90 mM NaCl, 1 mM EGTA, 1 mM DTT, 100 µM ATP, and 10 µM FMN, was added and incubated at 24 ℃ for 3 h in the dark or under blue light (100 µmol m-2 s-1). After washing the beads three times, the bound proteins were mixed with the SDS sample buffer and subjected to immunoblotting using anti-phot1 and anti-WDR48 antibodies.

For the in vitro pull-down assay using the N- and C-terminal fragments of phot1, FLAG-P1N or FLAG-P1C was diluted in buffer containing 20 mM Tris-HCl (pH 7.4), 140 mM NaCl, and 0.1% (v/v) Triton X-100, and incubated with anti-FLAG M2 affinity gel at 4 ℃ for 1 h. After washing the beads three times, His-WDR48, diluted in buffer containing 20 mM Tris-HCl (pH 7.4) and 140 mM NaCl, was added and incubated at 4 ℃ for 3 h in the dark. After washing the beads three times, the bound proteins were mixed with the SDS sample buffer and analysed by immunoblotting using anti-FLAG-HRP and anti-WDR48 antibodies.

In vitro kinase assay
His-phot1, His-D806N, His-phot2, and His-D720N were synthesised using an in vitro transcription/translation kit66 (BioSieg) supplemented with 10 µM FMN. The FLAG-WDR48 was synthesised using the same kit without FMN. His-tagged and FLAG-tagged proteins were incubated in buffer containing 37.5 mM Tris-HCl (pH 7.5), 5.3 mM MgSO , 90 mM NaCl, 1 mM EGTA, 1 mM DTT, 100 µM ATP, and 10 µM FMN at 24 ℃ for 1 h in the dark or under blue light (100 µmol m-2 s-1). For λ-PPase treatment, FLAG-WDR48 was recovered from the reaction mixture using anti-FLAG M2 affinity gel and washed three times. The beads were then incubated with λ-PPase in buffer containing 50 mM HEPES (pH 7.5), 100 mM NaCl, 2 mM DTT, 0.01% (w/v) Brij 35, and 1 mM MnCl at 30 ℃ for 1 h. The reaction mixtures were subjected to SDS-PAGE and analysed by immunoblotting with anti-pSer393 of WDR48, anti-WDR48, anti-phot1, and anti-phot2 antibodies.

Co-immunoprecipitation
Three-day-old etiolated seedlings expressing 3×FLAG-WDR48, as well as wild-type seedlings, were kept in the dark or illuminated with blue light (100 µmol m-2 s-1) for 2 min and then homogenised in extraction buffer containing 50 mM MOPS-KOH (pH 7.5), 2.5 mM EDTA, 100 mM NaCl, 0.5 mM PMSF, 10 µM leupeptin, 2 mM DTT, 10 mM NaF, 0.5 mM ammonium molybdate, and 100 nM calyclin A. After initial centrifugation at 10,000 × g for 10 min at 4 ℃, the supernatants were centrifuged at 100,000 × g for 1 h at 4 ℃ to obtain microsomal membrane. The membrane was resuspended in an extraction buffer containing 0.1% (v/v) Triton X-100 and kept on ice for 10 min. Following centrifugation at 10,000 × g for 10 min, the supernatant was subjected to immunoprecipitation with anti-FLAG M2 affinity gel for 12 h at 4 ℃. After washing the beads three times, the immunoprecipitated proteins were mixed with the SDS sample buffer and analysed by immunoblotting with anti-phot1 and anti-FLAG-HRP antibodies.

Subcellular localisation
Leaves from 2-week-old dark-adapted plants expressing phot1-GFP, GFP-WDR48, GFP-S393A, GFP-S393E, or co-expressing phot1-GFP and OEP7-mCherry or GFP-WDR48 and OEP7-mCherry, were illuminated with blue light (10 µmol m-2 s-1) for 10 min. GFP, mCherry, and chlorophyll fluorescence were then detected using a confocal laser scanning microscope (TCS SP8X; Leica Microsystems). For detection of GFP and mCherry fluorescence, the time gating method was applied with a gating time at 0.5–12 ns to eliminate background autofluorescence68. The excitation and emission wavelengths were 485 nm and 490–550 nm for GFP, 595 nm and 600–640 nm for mCherry, and 485 nm and 620–685 nm for chlorophyll fluorescence. The GFP signal at the outer edge of the guard cell was defined as the plasma membrane, whereas the signal inside the guard cell was defined as the cytosol. Fluorescence intensities in these regions were quantified using ImageJ 1.53q software (National Institutes of Health), and the cytosol/plasma membrane fluorescence intensity ratio was then calculated.

BiFC assay
The BiFC assay was performed according to a previously described method, with slight modifications27. The WDR48 cDNA sequence was amplified using primers 5′-TAGTGGATCCGTCGACATGCTACCGGGTTGGAAGTGC-3′ and 5′-TACCCTCGAGGTCGACTTATCTAGCTATAGCCACTC-3′, and the product was inserted downstream of the SPYNE(R) 173 sequence at the Sal  site. PHOT1 and PIN1 cDNA sequences were amplified using the following primers: 5′-TAGTGGATCCGTCGACATGGAACCAACAGAAAAACCATCGAC-3′ and 5′-TACCCTCGAGGTCGACTCAAAAAACATTTGTTTGCAGATCTTC-3′ for PHOT1, and 5′-TAGTGGATCCGTCGACATGATTACGGCGGCGGACTTC-3′ and 5′-TACCCTCGAGGTCGACTCATAGACCCAAGAGAATGTAG-3′ for PIN1. The resulting products were inserted downstream of the SPYCE (MR) sequence using the Sal  site. To construct the p35S:KAT1-mCherry vector, the mCherry sequence was inserted into the Sal /BamH  sites of pPCS201, generating pPCS205. The genomic sequence of KAT1 was amplified using primers 5′-ACATATGCCCGTCGACATGTCGATCTCTTGGACTCGAAATTTC-3′ and 5′-TGCTCACCATGTCGACATTTGATGAAAAATACAAATGATCACCATC-3′, and the product was inserted into the Sal  site of pPCS205. To construct the p35S:OEP7-mCherry vector, the cDNA sequences of OEP7 and mCherry were amplified using the following primers: 5′-ACATATGCCCGTCGACATGGGAAAAACTTCGGGAGCG-3′ and 5′-GCCCTTGCTCACCATGTCTTTGGTTGGGTCAGATTTG-3′ for OEP7, and 5′-ATGGTGAGCAAGGGCGAGGAG-3′ and 5′-TTCAGAATTCGGATCCTTACTTGTACAGCTCGTCCATG-3′ for mCherry. The products were inserted into the Sal /BamH  sites of pRI 101-AN. To construct the p35S:3×FLAG-WDR48 vector, the 3×FLAG sequence was inserted into the Sal /BamH  sites of pRI 101-AN to yield pPCS106. The WDR48 cDNA sequence was amplified using primers 5′-CGATGACAAGGGATCCATGCACCGGGTTGGAAGTGC-3′ and 5′-TTCAGAATTCGGATCCTTATCTAGCTATAGCCACTCTGTAGTTG-3′, and the product was inserted into the BamH  site of pPCS106.

The resulting vectors were transformed into mesophyll cell protoplasts using PEG-calcium transfection69. The adaxial epidermis of leaves from 4-week-old plants was removed, and the leaves were incubated in an enzyme solution containing 20 mM MES-KOH (pH 5.4), 0.25 M mannitol, 1% (w/v) cellulase R-10, 0.25% (w/v) macerozyme R-10, 0.1% (w/v) bovine serum albumin, 10 mM CaCl , and 20 mM KCl for 1 h. Mesophyll cell protoplasts were filtered through a 58 µm nylon mesh and collected by centrifugation at 100 × g for 3 min. The protoplasts were resuspended in W5 solution containing 2 mM MES-KOH (pH 5.7), 5 mM glucose, 154 mM NaCl, 125 mM CaCl , and 5 mM KCl and washed twice with W5 solution. The protoplasts were then resuspended in a buffer containing 4 mM MES-KOH (pH 5.7), 0.4 M mannitol, and 15 mM MgCl and mixed with 5 µg of each vector and 25 µg of competitor vector. Transformation was initiated by adding an equivalent volume of PEG solution containing 40% (w/v) polyethylene glycol 4,000, 0.2 M mannitol, and 100 mM CaCl . The reaction was halted by adding W5 solution, followed by centrifugation at 1,000 × g for 1 min. The protoplasts were resuspended in W5 solution and incubated for 24 h in the dark. YFP and mCherry fluorescence were detected using a confocal laser scanning microscope (TCS SP8X; Leica Microsystems) with a time-gating window of 0.5–12 ns to eliminate background autofluorescence68. The excitation and emission wavelengths were 514 nm and 520–550 nm for YFP, and 595 nm and 600–640 nm for mCherry. YFP and mCherry signals were quantified using ImageJ 1.53q software (National Institutes of Health), and the relative YFP fluorescence intensity was calculated based on the mCherry signal.

Chloroplast movements
Chloroplast movements were examined using green and white band assays, following previously described methods with slight modifications39. Leaves from dark-adapted plants were placed on 1% (w/v) agar plate, and slotted aluminum sheets were placed on top of the leaves. For the green band assay, leaves were illuminated with blue light (0.5 µmol m-2 s-1) for 1 h. For the white band assay, leaves were illuminated with blue light (50 µmol m-2 s-1) for 1 h.

Measurement of lateral root number and root length
Plants were grown for 11 days on 0.8% (w/v) agar plates containing 1× Murashige–Skoog (MS) salts, 25 mM MES-NaOH (pH 5.7), and 0.5% (w/v) sucrose under continuous white light (50 µmol m-2 s-1) at 23 ℃, as described previously36. Total lateral root number and root length were measured using ImageJ 1.53q software (National Institutes of Health).

Statistical analysis
Reported data are presented as mean ± the standard error (SEM). Significant differences were determined by analysis of variance (ANOVA) with Tukey’s test or Student’s t-test using Microsoft Excel and Excel Toukei ver. 6.05 and 8.0 (Esumi).

Peer review rounds
[{"round": 1, "reviewer_comments": {"reviewer_1": "In this manuscript, the authors investigate the novel function of WDR48 in the blue light-mediated guard cell starch degradation and stomatal opening. Through phosphoproteome analysis, the authors identified WDR48 as a protein that is phosphorylated when exposed to blue light. Furthermore, PHOT1 was found to interact with and phosphorylate WDR48 at Ser 393. WDR48 and BLUS1 regulate guard cell starch degradation and stomatal opening through distinct pathways. The data presented in the manuscript is original and will be of interest to a wide range of researchers studying light signaling and stomatal movement. Some of the findings could be further supported by addressing the following concerns:\n1. The authors conducted the phosphoproteomic analysis using guard cell protoplasts from wild type and phot1-5 phot2-1 mutants. However, it is unclear which materials were used in Figure 1a and 1b, and this information should be included in the figure legend. Furthermore, the manuscript does not indicate how many times these experiments were repeated and it will be better to perform the statistical analysis.\n2. Blue light induces the phosphorylation of WDR48 via PHOT1/PHOT2, which promotes starch degradation in guard cells. It is unclear whether mimicking the phosphorylation of WDR48-S393E could alleviate the defective phenotype of impaired guard cell starch degradation and stomatal opening observed in phot1 phot2 mutants. In other words, as a downstream component of the PHOT1/PHOT2-mediated blue light signaling pathway, activated WDR48 should repress the defective phenotype of phot1phot2.\n3. What is the biological significance of PHOT1-dependent phosphorylation modification of WDR48 protein, and how does this affect the protein localization of WDR48-S393A or D? Does this phosphorylation modification influence the binding ability of WDR48 to deubiquitinating enzymes (DUBs)?\n4. It has been reported that WDR48 regulates lateral root initiation. It is unclear whether the blue light-induced phosphorylation of WDR48 has any effect on the regulation of lateral root growth by WDR48.\n5. In Figure 5a, the confocal image depicts PHOT1-GFP localized in the plasma membrane, whereas GFP-WDR48 is found both in the plasma membrane and cytoplasm. It would be beneficial to perform a statistical analysis to determine whether blue light specifically regulates the subcellular localization of GFP-WDR48.\n6. The BiFC experiments indicate that both PHOT1 and WDR48 are localized in the chloroplast, and PHOT1/PHOT2 play critical roles in regulating chloroplast movement in response to light. It is unclear whether WDR48 is also involved in this process. Additionally, it would be helpful if Figure 5a included an image depicting the chloroplast localization of PHOT1 and WDR48.\n7. The observation of blue light-induced phosphorylation of WDR48 in the aha1-9 mutant suggests that H+-ATPase is not required for this process. However, this data does not support the hypothesis that WDR48 regulates guard cell starch degradation through an H+-ATPase-independent pathway. It would be interesting to investigate whether activated WDR48-S393E can alleviate the defective starch degradation phenotype observed in aha1-9, blus1, and cbc1 mutants.\n8. All gel images should include the molecular weight marker and original image, and the relative quantified protein band intensity compared to the non-phosphorylated protein should be presented.", "reviewer_2": "The current manuscript aims to explore the contributions of WDR48 in mediating the starch degradation and stomatal opening via the phosphorylation of blue light irradiated Phototropins. The authors identified WDR48 as a phosphorylated protein using mass spectrometry analysis under blue light conditions. By employing a mix of biochemical assays, PS-PI staining, confocal imaging, and immunoprecipitation, the authors propose that the phosphorylation of WDR48 is associated with blue light-induced starch degradation and stomatal opening. Additionally, this study establishes that WDR48 is phosphorylated by Phototropins. This work also presents other interesting claims, including the separate signaling pathways of WDR48 and BLUS1, which coordinately regulate starch degradation and synthesis. However, weaknesses in the data must be addressed before these claims can be fully supported.\nThere are several major limitations to this work:\n1- The majority of data regarding the phosphorylation of WDR48 was obtained solely through the use of phospho-specific antibodies against Ser-393 of WDR-48. Considering the phosphorylation of WDR48 is a central claim in this paper and the authors have access to the mass spec facilities, presenting mass spectrometry data to support the key phosphorylation results presented via Western blot would be highly beneficial.\nSecondly, based on mass spec data (Fig. 1b), the phosphorylation of WDR48 in blue light is noticeably stronger than that in red light (the peak of phosphopeptide containing\npSer-393 of WDR48 is almost undetectable in RL compare to that of in BL). However,\nin most of WB assay (e.g. Fig. 1d), the band of WDR48 is clearly presented in RL of WT and in phot1-5/phot2-1. The authors should address the discrepancy between the mass spec and WB data, or consider the possibility that the antibody targeting Ser-393 may detect a non-specific protein with similar electrophoretic mobility to phosphorylated WDR48 (if this is the case, additional mass spec data for WDR48 phosphorylation should be presented).\n2-The resolution of all the figures should be significantly improved. The current quality of figures on this MS does not meet the standards expected for publication in Nature Communications IMO. Most starch granules stained with PS-PI are hard to distinguish in this MS, especially those undergoing degradation. The authors could refer to ref.#28 to modify their figure (the current biology paper:” Blue light induces a distinct starch degradation pathway in guard cells for stomatal opening.”).\n3-To examine whether WDR48 interacts with phototropins, the authors detected the interaction of WDR48 and phototropins by in vitro pull down, co-immunoprecipitation and BiFC assay. However, the input data of the bait were missing in the pull down (Fig. 4f) and co-immunoprecipitation (Fig. 4g) assay. To me, the similar amount of prey (input of bait) is also important to assess the protein-protein interaction.\nThe Lines 202—204: The authors mention that“In the dark, GFP fluorescence of PHOT1-GFP was detected at the plasma membrane, and partial GFP signals were detected in the cytoplasm after blue light irradiation (Fig. 5a). ” Regarding the BiFC assay, the authors should specify the light conditions under which it was conducted, considering that no PHOT1-GFP signal was detected in the chloroplast envelope in the dark.\nSince the interaction and phosphorylation of WDR48 are the main claims of this paper, the authors should illustrate whether WDR48 interacts with and is phosphorylate by phototropins in dark via pull down, co-immunoprecipitation and in vitro kinase assay, which could support the phosphorylation data detected by antibodies against Ser-393 of WDR-48.\nOther comments:\n(1) The Confocal images of PS-PI staining (Fig. 2e) do not match the Quantification data (Fig. 2f). e.g. the starch granule of phot1-5/phot2-1 at 5 and 10 min.\n(2) The trend of changes in stomatal aperture (Fig. 2g) is inconsistent with the curve in Fig. 2e. A confocal image depicting a consistent trend can be acquired to complement Fig. 2g.\n(3) #3 line of GFP-S393A exhibits a discernible disparity from #9. Did the authors examine the protein expression levels of both lines?\n(4) The Confocal images of Fig. 3a do not correspond to the statistical findings (both starch granule area and stomatal aperture) reported by the authors. Alternatively, it is possible that my interpretation was influenced by the poor quality of the images.\n(5) In Figure 4, it can be observed that BLUS1 protein (using anti-BLUS1) exhibits distinct shifted bands indicative of phosphorylation after blue light irradiation. However, such shifted bands are not visible in the case of WDR48 (using anti-WDR48), and the phosphorylation of WDR48 can only be detected using anti-pSer393. This is one of my concerns regarding the phosphorylation detection in this study.\n(6) The phosphorylation of WDR48 in the in vitro kinase assay (Fig. 4e) should be confirmed by λ-PPase.\n(7) The author should explain that, in Fig. 4b, why the band related to anti-pSer393 was not detectable in WT under RL, compared to the same situation in other figures, e.g. Fig. 4a.\n(8) Since phot1-GFP refers to a protein, it should be written as PHOT1-GFP."}, "author_response": "Response to Reviewers\nReviewer #1 (Remarks to the Author):\nIn this manuscript, the authors investigate the novel function of WDR48 in the blue light-mediated\nguard cell starch degradation and stomatal opening. Through phosphoproteome analysis, the authors\nidentified WDR48 as a protein that is phosphorylated when exposed to blue light. Furthermore,\nPHOT1 was found to interact with and phosphorylate WDR48 at Ser 393. WDR48 and BLUS1\nregulate guard cell starch degradation and stomatal opening through distinct pathways. The data\npresented in the manuscript is original and will be of interest to a wide range of researchers studying\nlight signaling and stomatal movement. Some of the findings could be further supported by addressing\nthe following concerns:\n1. The authors conducted the phosphoproteomic analysis using guard cell protoplasts from wild type\nand phot1-5 phot2-1 mutants. However, it is unclear which materials were used in Figure 1a and 1b,\nand this information should be included in the figure legend. Furthermore, the manuscript does not\nindicate how many times these experiments were repeated and it will be better to perform the\nstatistical analysis.\nResponse: Thank you for your suggestions. We have added the information of the materials used for\nthe phosphoproteome analysis to the figure legend of Fig. 1a and 1b. We also provided the relative\npeak area of the phosphopeptide containing pSer-393 of WDR48 in the wild type and the phot1 phot2\nmutant, along with the number of replicates and the results of statistical analysis.\n2. Blue light induces the phosphorylation of WDR48 via PHOT1/PHOT2, which promotes starch\ndegradation in guard cells. It is unclear whether mimicking the phosphorylation of WDR48-S393E\ncould alleviate the defective phenotype of impaired guard cell starch degradation and stomatal opening\nobserved in phot1 phot2 mutants. In other words, as a downstream component of the PHOT1/PHOT2-\nmediated blue light signaling pathway, activated WDR48 should repress the defective phenotype of\nphot1phot2.\nResponse: Thank you for your insightful comment. To address this point, we crossed the transgenic\nplants expressing GFP-S393E (generated in the wdr48-1 mutant background) with the phot1-5 phot2-\n1 double mutant, thereby obtaining plants expressing GFP-S393E in the phot1-5 phot2-1 wdr48-1\ntriple-mutant background. We then examined starch degradation and stomatal opening in response\nto blue light and fusicoccin, a plasma membrane H⁺-ATPase activator (Fig. 6g–i).\nIn these plants, we found that blue light-induced starch degradation and stomatal opening were not\nrestored, indicating that the phosphomimetic S393E mutation alone is insufficient to rescue these\ndefects in the absence of phot1 and phot2. In contrast, fusicoccin treatment successfully induced starch\ndegradation and stomatal opening, even without blue light irradiation.\nThese observations suggest the following interpretation. Although the S393E substitution likely\nrenders WDR48 constitutively active in a manner similar to its phosphorylated state, the loss of phot1\nand phot2 prevents blue light-dependent phosphorylation of BLUS1, a key upstream regulator\nrequired for activation of the plasma membrane H⁺-ATPase. As a result, the plasma membrane\nhyperpolarisation and the downstream starch degradation pathway, which depends on H⁺-ATPase\nactivity, are all impaired in the phot1-5 phot2-1 background. This explains why blue light-induced\nstarch degradation and stomatal opening does not occur even when WDR48 is constitutively activated.\nIn contrast, fusicoccin bypasses the phototropin–BLUS1 pathway by directly activating H⁺-\nATPase, thereby restoring plasma membrane hyperpolarisation and initiating the downstream\nsignalling that leads to starch degradation. Additionally, because WDR48 is already activated by the\nS393E mutation, both signalling branches are functional under fusicoccin treatment, resulting in the\nobserved induction of starch degradation and stomatal opening.\nTaken together, these findings indicate that although the phosphorylation-mimicking activation of\nWDR48 is important, it is not sufficient to compensate for the loss of the phototropin–BLUS1–H⁺-\nATPase signalling cascade, which is essential for blue light-induced guard cell responses.\n3. What is the biological significance of PHOT1-dependent phosphorylation modification of WDR48\nprotein, and how does this affect the protein localization of WDR48-S393A or D? Does this\nphosphorylation modification influence the binding ability of WDR48 to deubiquitinating enzymes\n(DUBs)?\nResponse: Thank you for your insightful questions. At present, the biological significance of\nphototropin-mediated phosphorylation of WDR48 remains unclear. We have conducted additional\nanalyses on the subcellular localisation of GFP-S393A and GFP-S393E, but no noticeable differences\nwere observed compared with GFP-WDR48, indicating that this phosphorylation modification does\nnot affect WDR48 localisation (Supplementary Fig. 8).\nRegarding the potential influence of WDR48 phosphorylation on its interaction with\ndeubiquitinating enzymes (DUBs), we acknowledge that such an effect is conceivable. However,\ninvestigating this question is beyond the scope of the current study, as it is not yet established whether\nWDR48 interacts with DUBs in guard cells or whether DUBs contribute to starch degradation and\nstomatal opening. We consider this an interesting topic for future research.\n4. It has been reported that WDR48 regulates lateral root initiation. It is unclear whether the blue light-\ninduced phosphorylation of WDR48 has any effect on the regulation of lateral root growth by WDR48.\nResponse: Thank you for your insightful comment. We indeed confirmed not only the inhibition of\nlateral root formation but also the suppression of primary root growth in the wdr48 mutants\n(Supplementary Fig. 5). However, these phenotypes were complemented by the introduction of wild-\ntype WDR48 as well as by the phosphodefective and phosphomimetic forms of WDR48. Thus, it\nappears that these root developmental processes require functions of WDR48 that are independent of\nits phosphorylation status. We have incorporated these findings into the Results section.\n5. In Figure 5a, the confocal image depicts PHOT1-GFP localized in the plasma membrane, whereas\nGFP-WDR48 is found both in the plasma membrane and cytoplasm. It would be beneficial to perform\na statistical analysis to determine whether blue light specifically regulates the subcellular localization\nof GFP-WDR48.\nResponse: Thank you for this valuable suggestion. In response to your comment, we performed a\nquantitative analysis of fluorescence intensity in both the plasma membrane and cytosolic fractions,\nand evaluated their relative distribution using the cytosol/plasma membrane ratio for GFP-WDR48 as\nwell as for phot1-GFP (Fig. 5a and 5b).\nOur statistical analysis revealed that phot1-GFP exhibited a significant increase in the\ncytosol/plasma membrane ratio upon blue light irradiation, consistent with the partial release of phot\n1 from the plasma membrane into the cytosol under blue light conditions (Sakamoto and Briggs, 2002;\nWan et al., 2008). In contrast, GFP-WDR48 did not show any detectable change in the cytosol/plasma\nmembrane ratio following blue light exposure, indicating that its subcellular localisation remains\nstable during the blue light response.\nThese results demonstrate that, unlike phot1, the localisation of WDR48 is not dynamically\nregulated by blue light within the timescale examined. We have incorporated these new quantitative\ndata into the revised manuscript (Fig. 5b).\n(Reference)\nSakamoto and Briggs (2002) Plant Cell 14: 1723–1735.\nWan et al. (2008) Mol Plant 1: 103–117.\n6. The BiFC experiments indicate that both PHOT1 and WDR48 are localized in the chloroplast, and\nPHOT1/PHOT2 play critical roles in regulating chloroplast movement in response to light. It is unclear\nwhether WDR48 is also involved in this process. Additionally, it would be helpful if Figure 5a included\nan image depicting the chloroplast localization of PHOT1 and WDR48.\nResponse: We appreciate the reviewer’s insightful comments regarding the potential involvement of\nWDR48 in chloroplast movements and the localisation of phot1 and WDR48 on the chloroplast\nenvelope. To address these points, we performed additional analyses using the wdr48 mutants. Our\nresults demonstrate that both the weak blue light-induced chloroplast accumulation and the strong blue\nlight-induced chloroplast avoidance occur normally in the wdr48 mutants, comparable to those in the\nwild type (Supplementary Fig. 3). These findings indicate that WDR48 is not essential for\nchloroplast movements.\nIn addition, following the reviewer’s suggestion regarding Figure 5a, we conducted additional\nanalyses to examine the chloroplast envelope localisation of phot1-GFP and GFP-WDR48 in more\ndetail. We introduced the chloroplast envelope marker OEP7-mCherry into plants expressing phot1-\nGFP or GFP-WDR48 and confirmed that the GFP signals of both fusion proteins colocalised with\nOEP7-mCherry under both dark and blue light conditions. These findings indicate that both phot1 and\nWDR48 are partially localised to the chloroplast envelope irrespective of light conditions.\nWe have included representative images illustrating their colocalisation on the chloroplast envelope,\nas well as images depicting the chloroplast region, which now provided as Supplementary Figs. 9a\nand 9b, respectively.\n7. The observation of blue light-induced phosphorylation of WDR48 in the aha1-9 mutant suggests\nthat H+-ATPase is not required for this process. However, this data does not support the hypothesis\nthat WDR48 regulates guard cell starch degradation through an H+-ATPase-independent pathway. It\nwould be interesting to investigate whether activated WDR48-S393E can alleviate the defective starch\ndegradation phenotype observed in aha1-9, blus1, and cbc1 mutants.\nResponse: Thank you for this insightful comment. Following our analysis in the phot1-5 phot2-\n1 background described above, we conducted similar experiments to evaluate whether activated\nWDR48-S393E can alleviate the defects in guard cell starch degradation observed in blus1-\n1 and aha1-9 mutants.\nTo address this, we crossed the wdr48-1 expressing GFP-S393E with blus1-1 and aha1-9 mutants,\nand obtained plants expressing GFP-S393E in the blus1-1 wdr48-1 and aha1-9 wdr48-1 mutant\nbackgrounds. We then examined starch degradation and stomatal opening in response to blue light\nand fusicoccin (Fig. 6g–i).\nIn the blus1-1 wdr48-1 background expressing GFP-S393E, blue light did not induce starch\ndegradation or stomatal opening, consistent with the results in the phot1-5 phot2-1 wdr48-1\nbackground described above. However, fusicoccin treatment successfully induced both starch\ndegradation and stomatal opening, indicating that although S393E is constitutively active, BLUS1-\ndependent H⁺-ATPase activation remains essential for the blue light-triggered response.\nIn contrast, in the aha1-9 wdr48-1 background expressing GFP-S393E, neither blue light nor\nfusicoccin triggered starch degradation or stomatal opening. Since the aha1-9 mutant exhibits reduced\nexpression of the H⁺-ATPase, it is conceivable that even fusicoccin is unable to sufficiently induce\nplasma membrane hyperpolarisation and downstream signalling required for starch degradation. This\nexplains why GFP-S393E cannot compensate for the defective phenotype in this background.\nAlthough the data are not shown, blue light-dependent starch degradation in the cbc1-1 cbc2-\n1 double mutant occurred normally, similar to the wild type. Therefore, we did not conduct further\nanalyses using plants expressing GFP-S393E in the cbc1-1 cbc2-1 wdr48-1 mutant background.\nTaken together, these results collectively suggest that although the phosphomimetic S393E\nmutation activates WDR48, its ability to promote starch degradation still depends on a functional level\nof H⁺-ATPase activity. Thus, WDR48 alone is insufficient to drive starch degradation when the H⁺-\nATPase activation is severely compromised, as in the blus1-1 and aha1-9 mutants. The results\ndescribed above, including those obtained using GFP-S393E in the phot1 phot2 wdr48 mutant\nbackground, have been incorporated into the revised manuscript, and the implications of these findings\nare thoroughly discussed in the Discussion section.\n8. All gel images should include the molecular weight marker and original image and the relative\nquantified protein band intensity compared to the non-phosphorylated protein should be presented.\nResponse: Thank you for your comments. We have included the molecular weight information in all\ngel images (Figs. 1d, 1g, 1k, 1m, 1n, 4a, 4c, 4e, 4g, 4i, 4k, and 4m; Supplementary Fig. 1a, 1c, 1e, 4a;\n7a, 7c, 7e, 10a, and 10c), and we have provided the original gel images in the Source Data. In addition,\nwe have provided the quantitative data for the relative phosphorylation levels (Figs. 1e, 1l, 4b, 4d, 4f,\nand 4h; Supplementary Fig. 1b, 1d, 1f, 7b, 7d, 7f, 10b, and 10d).\nReviewer #2 (Remarks to the Author):\nThe current manuscript aims to explore the contributions of WDR48 in mediating the starch\ndegradation and stomatal opening via the phosphorylation of blue light irradiated Phototropins. The\nauthors identified WDR48 as a phosphorylated protein using mass spectrometry analysis under blue\nlight conditions. By employing a mix of biochemical assays, PS-PI staining, confocal imaging, and\nimmunoprecipitation, the authors propose that the phosphorylation of WDR48 is associated with blue\nlight-induced starch degradation and stomatal opening. Additionally, this study establishes that\nWDR48 is phosphorylated by Phototropins. This work also presents other interesting claims, including\nthe separate signaling pathways of WDR48 and BLUS1, which coordinately regulate starch\ndegradation and synthesis. However, weaknesses in the data must be addressed before these claims\ncan be fully supported.\nThere are several major limitations to this work:\n1- The majority of data regarding the phosphorylation of WDR48 was obtained solely through the use\nof phospho-specific antibodies against Ser-393 of WDR-48. Considering the phosphorylation of\nWDR48 is a central claim in this paper and the authors have access to the mass spec facilities,\npresenting mass spectrometry data to support the key phosphorylation results presented via Western\nblot would be highly beneficial. Secondly, based on mass spec data (Fig. 1b), the phosphorylation of\nWDR48 in blue light is noticeably stronger than that in red light (the peak of phosphopeptide\ncontaining pSer-393 of WDR48 is almost undetectable in RL compare to that of in BL). However, in\nmost of WB assay (e.g. Fig. 1d), the band of WDR48 is clearly presented in RL of WT and in phot1-\n5/phot2-1. The authors should address the discrepancy between the mass spec and WB data, or\nconsider the possibility that the antibody targeting Ser-393 may detect a non-specific protein with\nsimilar electrophoretic mobility to phosphorylated WDR48 (if this is the case, additional mass spec\ndata for WDR48 phosphorylation should be presented).\nResponse: We thank the reviewer for the insightful comments regarding the phosphorylation of\nWDR48. Initially, we performed phosphoproteomic analysis using guard cell protoplasts isolated from\nArabidopsis wild type, and identified WDR48 as a blue light-dependent phosphorylated protein. As\nguard cell protoplasts can only be obtained in small amounts from a large quantity of plant leaves, it\nis not feasible to perform phosphoproteomic analysis for all biochemical experiments. Therefore, we\ngenerated phospho-specific antibodies recognizing WDR48 phosphorylated at Ser-393 and used them\nto assess WDR48 phosphorylation under various conditions and in different mutant backgrounds.\nTo address the reviewer’s concerns, we reperformed phosphoproteomic analysis using guard cell\nprotoplasts from both wild type and phot1-5 phot2-1 mutant. Quantitative analysis of the Ser-393-\ncontaining phosphopeptide revealed that this phosphopeptide is indeed detectable under red light in\nthe wild type, and its abundance increases by approximately 3.5-fold upon blue light irradiation. In\nthe phot1-5 phot2-1 mutant, the phosphopeptide is detected under red light at a level similar to that of\nwild-type red light samples; however, blue light irradiation does not increase its abundance. These\nquantitative values have been added to Fig. 1b.\nWe also conducted a detailed validation of the specificity of the phospho-specific antibodies against\nSer-393 of WDR48 used in our biochemical analyses. In wild-type guard cell protoplasts, a weak\nsignal was detectable even under red light, and the signal increased upon blue light irradiation. λ-\nprotein phosphatase treatment reduced the signal to below the detection limit under both red and blue\nlight conditions (Supplementary Figs. 1a and 1b). In the wdr48 null mutant, no signal was detected at\nthe molecular weight corresponding to WDR48, ruling out cross-reactivity with other proteins of\nsimilar size (Supplementary Figs. 1c and 1d). Furthermore, in the phosphodefective WDR48 mutant\n(S393A), the antibodies did not detect any band at the WDR48 molecular weight (Supplementary Figs.\n1e and 1f). These results confirm that the antibodies specifically recognise Ser-393-phosphorylated\nWDR48.\nFinally, we performed quantitative analysis for all biochemical assays using these antibodies (Fig.\n1e, 4b, 4d, 4f, and 4h; Supplementary Figs. 1b, 1d, 1f, 7b, 7d, 7f, 10b, and 10d). As an example, in\nwild type and the phot1-5 phot2-1 mutant, the relative phosphorylation level (with wild-type blue light\nset as 100%) showed approximately 20% signal under red light for both genotypes, and no increase\nwas observed in phot1-5 phot2-1 plants under blue light (Fig. 1e). These results are consistent with\nthe quantitative data from phosphoproteomic analysis (Fig. 1b).\nTaken together, these results demonstrate that WDR48 undergoes low-level basal phosphorylation\nunder red light in guard cells, that its phosphorylation levels are strongly enhanced by blue light,\nand that this blue light-dependent phosphorylation is dependent on phototropins.\n2-The resolution of all the figures should be significantly improved. The current quality of figures on\nthis MS does not meet the standards expected for publication in Nature Communications IMO. Most\nstarch granules stained with PS-PI are hard to distinguish in this MS, especially those undergoing\ndegradation. The authors could refer to ref.#28 to modify their figure (the current biology paper:” Blue\nlight induces a distinct starch degradation pathway in guard cells for stomatal opening.”).\nResponse: Thank you for your valuable comment. We have carefully reexamined the starch granule\ndata and found that some of the original images did not clearly represent the degradation status of\nstarch granules in guard cells. To address this, we have replaced the images with higher resolution\nconfocal images that more clearly depict starch granule morphology and degradation. We believe these\nupdated images improve the visual clarity and accurately correspond to the quantified data, ensuring\nthat the figures meet the standards expected for publication.\n3-To examine whether WDR48 interacts with phototropins, the authors detected the interaction of\nWDR48 and phototropins by in vitro pull down, co-immunoprecipitation and BiFC assay. However,\nthe input data of the bait were missing in the pull down (Fig. 4f) and co-immunoprecipitation (Fig. 4g)\nassay. To me, the similar amount of prey (input of bait) is also important to assess the protein-protein\ninteraction.\nResponse: Thank you for your comment. In response, we have repeated the in vitro pull-down and co-\nimmunoprecipitation assays and included the input data for both the prey and the bait (Figs. 4i, 4k,\nand 4m). The reanalysed results confirm the interaction between phototropins and WDR48, consistent\nwith our original findings.\nThe Lines 202—204: The authors mention that “In the dark, GFP fluorescence of PHOT1-GFP was\ndetected at the plasma membrane, and partial GFP signals were detected in the cytoplasm after blue\nlight irradiation (Fig. 5a). ” Regarding the BiFC assay, the authors should specify the light conditions\nunder which it was conducted, considering that no PHOT1-GFP signal was detected in the chloroplast\nenvelope in the dark.\nResponse: We appreciate the reviewer’s insightful comment regarding the light conditions used for\nthe BiFC assay. The BiFC experiments were performed using dark-treated protoplasts. During our\nobservations, we noted that in the dark, phot1-GFP fluorescence is mainly observed at the plasma\nmembrane, with weaker signals occasionally detectable around chloroplasts, and that blue light\nirradiation enhances its cytoplasmic localisation, as shown in Fig. 5a.\nTo clarify its localisation on the chloroplast envelope more precisely, we introduced the chloroplast\nenvelope marker OEP7-mCherry into plants expressing phot1-GFP, as well as those expressing GFP-\nWDR48, and performed additional analyses (Supplementary Fig. 9). These analyses revealed that\nphot1-GFP signals are detectable on the chloroplast envelope under both dark and blue light conditions,\nconsistent with a previous report (Kong et al., 2013), which was confirmed by colocalisation with\nOEP7-mCherry. Similarly, GFP-WDR48 also localises to the chloroplast envelope in both conditions.\nThese findings explain why BiFC signals appear on the chloroplast envelope even in dark-treated\nprotoplasts and address the reviewer’s concerns regarding phot1-GFP localisation. We have revised\nthe relevant section of the manuscript accordingly and incorporated these new data to more accurately\nreflect phot1-GFP and GFP-WDR48 behavior under dark and blue light conditions.\n(Reference)\nKong et al. (2013) Plant Cell Physiol 54: 80–92.\nSince the interaction and phosphorylation of WDR48 are the main claims of this paper, the authors\nshould illustrate whether WDR48 interacts with and is phosphorylate by phototropins in dark via pull\ndown, co-immunoprecipitation and in vitro kinase assay, which could support the phosphorylation\ndata detected by antibodies against Ser-393 of WDR-48.\nResponse: Thank you for your comment. According to your comment, we reanalysed the interaction\nbetween WDR48 and phototropins under both dark and blue light conditions using in vitro pull-down\nand co-immunoprecipitation assays. These analyses revealed that WDR48 binds to phototropins\nregardless of blue light exposure (Figs. 4i–l). Additionally, we performed in vitro kinase assays using\nfull-length recombinant phototropins and confirmed the blue light-dependent phosphorylation of\nWDR48 (Figs. 4g and 4h; Supplementary Figs. 7a and 7b), consistent with in vivo phosphorylation\ndata. These results further support our conclusion that WDR48 is phosphorylated by phototropins in a\nblue light-dependent manner.\nOther comments:\n(1) The Confocal images of PS-PI staining (Fig. 2e) do not match the Quantification data (Fig. 2f). e.g.\nthe starch granule of phot1-5/phot2-1 at 5 and 10 min.\nResponse: Thank you for your comment. We have replaced the images in Fig. 2e with the correct\nconfocal images that accurately correspond to the quantification data shown in Fig. 2f. This ensures\nconsistency between the visual representation of starch granules and the quantified measurements for\nall genotypes, including phot1-5 phot2-1 at 5 and 10 min.\n(2) The trend of changes in stomatal aperture (Fig. 2g) is inconsistent with the curve in Fig. 2e. A\nconfocal image depicting a consistent trend can be acquired to complement Fig. 2g.\nResponse: Thank you for your comment. Indeed, in the data for starch degradation shown in Fig. 2e\nand 2f, starch in the wild type is largely degraded within 5 min of blue light irradiation. In contrast,\nstomatal opening (Fig. 2g) is initiated at 5 min but continues to increase until 10 min, showing a\ndifferent time course.\nThis difference likely reflects the physiological process linking starch degradation to stomatal\nopening: as the degradation products must be converted into osmolytes such as glucose and malate, a\nprocess that requires time. Glucose functions as an osmotic regulator, while malate acts as a counter-\nion for K+ (Ogawa et al., 1978; Shimazaki et al., 2007; Dong et al., 2018; Flütsch et al. 2020). Together,\nthese products drive water influx into guard cells, resulting in stomatal opening. Consequently, a\ntemporal lag between the rapid starch degradation and the subsequent stomatal opening is expected.\nTherefore, the observed difference in time course between starch degradation and stomatal opening is\nconsistent with the underlying physiological mechanism.\n(Reference)\nOgawa et al. (1978) Planta 142: 61–65.\nShimazaki et al. (2007) Annu Rev Plant Biol 58: 219–247.\nDong et al. (2018) Mol Plant 11: 1278–1291.\nFlütsch et al. (2020) Plant Cell 32: 2325–2344.\n(3) #3 line of GFP-S393A exhibits a discernible disparity from #9. Did the authors examine the protein\nexpression levels of both lines?\nResponse: Thank you for your comment. There is no statistically significant difference between the\n#3 and #9 lines of GFP-S393A in terms of blue light-dependent starch degradation and stomatal\nopening (Fig. 3a–c). We also examined the protein expression levels of both lines and found no\nstatistically significant difference; both lines express GFP-S393A at levels comparable to the\nendogenous WDR48 in the wild type (Supplementary Figs. 4a and 4b).\n(4) The Confocal images of Fig. 3a do not correspond to the statistical findings (both starch granule area\nand stomatal aperture) reported by the authors. Alternatively, it is possible that my interpretation\nwas influenced by the poor quality of the images.\nResponse: Thank you for your comment. We have replaced the confocal images in Fig. 3a with higher-\nquality images that accurately correspond to the statistical findings for both starch granule area and\nstomatal aperture. This ensures that the visual representation is fully consistent with the quantified\ndata and provides a clearer depiction of the observed phenotypes.\n(5) In Figure 4, it can be observed that BLUS1 protein (using anti-BLUS1) exhibits distinct shifted\nbands indicative of phosphorylation after blue light irradiation. However, such shifted bands are not\nvisible in the case of WDR48 (using anti-WDR48), and the phosphorylation of WDR48 can only be\ndetected using anti-pSer393. This is one of my concerns regarding the phosphorylation detection in\nthis study.\nResponse: Thank you for your insightful comment. While BLUS1 shows a clear band shift upon blue-\nlight-dependent phosphorylation, WDR48 does not exhibit a detectable mobility shift in SDS-\nPAGE. Phosphorylation does not always result in a detectable mobility shift on SDS-PAGE. This\ndifference likely reflects intrinsic properties of WDR48. Phosphorylation of WDR48 does not appear\nto induce a conformational change sufficient to alter its electrophoretic mobility, and the amino acid\ncontext around the phosphorylation site, including local charge distribution, may further prevent a\nvisible shift.\nImportantly, the phosphorylation of WDR48 under blue light has been independently confirmed\nby phosphoproteomic analysis, in addition to detection using the phospho-specific antibodies (anti-\npSer393) and λ-phosphatase treatment. These results collectively demonstrate that WDR48 is indeed\nphosphorylated under blue light, even though no mobility shift is observed on SDS-PAGE.\n(6) The phosphorylation of WDR48 in the in vitro kinase assay (Fig. 4e) should be confirmed by λ-\nPPase.\nResponse: Thank you for your valuable comment. Following your suggestion, we performed an in\nvitro kinase assay using full-length phot1 and phot2 and confirmed the blue light-dependent\nphosphorylation of WDR48. We further verified that these phosphorylation signals were reduced by\nλ-PPase treatment. These results demonstrate that WDR48 is indeed phosphorylated by phot1 and\nphot2 in a blue light-dependent manner in vitro.\n(7) The author should explain that, in Fig. 4b, why the band related to anti-pSer393 was not detectable\nin WT under RL, compared to the same situation in other figures, e.g. Fig. 4a.\nResponse: Thank you for pointing this out. In Fig. 4b, the anti-pSer393 signal in the wild type under\nRL was not detectable because the blot shown in the original version had been captured with a shorter\nexposure time when compared to other figures such as Fig. 4a. To avoid any misunderstanding, we\nhave replaced the panel with a blot obtained using a longer exposure time, in which the band is clearly\nvisible.\n(8) Since phot1-GFP refers to a protein, it should be written as PHOT1-GFP.\nResponse: We understand your concern. According to the established nomenclature rules for\nphototropin proteins, the holoprotein containing the chromophore flavin mononucleotide (FMN) is\ndenoted as phot, whereas the apoprotein lacking FMN is written as PHOT. Because phototropin\ntranslated in vivo predominantly exists as the holoprotein form, we have designated the protein\nas phot1-GFP throughout the manuscript.\n(Reference)\nBriggs et al. (2001) Plant Cell 13: 993–997."}, {"round": 2, "reviewer_comments": {"reviewer_1": "In the revised manuscript, the authors have effectively addressed most of my concerns. The paper is now more comprehensive and supported by stronger data. I have a few minor suggestions to further enhance the quality of this manuscript.\n1. It would be beneficial to present a new working model that including more information.\n2. The images in Figure 5A seem inconsistent with those in Supplementary Figures 8 and 9, particularly regarding the GFP-WDR48 images. In Figure 5A, GFP-WDR48 appears relatively blurry and seems to predominantly localize to the cytoplasm, whereas in Supplementary Figures 8 and 9, GFP-WDR48 displays clearer localization at the plasma membrane and chloroplast envelope, similar to GFP-PHOT1.\n3. There are several spelling errors throughout the manuscript. For instance, \"recognise\" in line 118 should be corrected to \"recognize,\" and \"localisation\" in line 226 should be changed to \"localization.\"", "reviewer_2": "I consider my comments to have been fully addressed, and I have no further questions."}, "author_response": "RESPONSE TO REVIEWER COMMENTS\nReviewer #1 (Remarks to the Author):\nIn the revised manuscript, the authors have effectively addressed most of my concerns. The paper is\nnow more comprehensive and supported by stronger data. I have a few minor suggestions to further\nenhance the quality of this manuscript.\n1. It would be beneficial to present a new working model that including more information.\nWe thank the reviewer for the suggestion. In response, we have incorporated additional information\ninto a new working model, presented in Fig. 7. This updated model includes starch synthesis via\nphotosynthesis under red light and starch degradation mediated by phototropin-dependent\nphosphorylation of BLUS1 and WDR48 under blue light. It illustrates how red and blue light\nsynergistically promote stomatal opening.\n2. The images in Figure 5A seem inconsistent with those in Supplementary Figures 8 and 9,\nparticularly regarding the GFP-WDR48 images. In Figure 5A, GFP-WDR48 appears relatively blurry\nand seems to predominantly localize to the cytoplasm, whereas in Supplementary Figures 8 and 9,\nGFP-WDR48 displays clearer localization at the plasma membrane and chloroplast envelope, similar\nto GFP-PHOT1.\nWe thank the reviewer for this careful observation. Upon reviewing all imaging data, we found that\nthe confocal images of GFP-WDR48 shown in Fig. 5A did not capture the plasma membrane and\nchloroplast envelope signals well due to limitations in focal depth. We have now replaced these images\nwith confocal images taken at similar focal depths to those in Supplementary Fig. 9 and 10, which\nmore accurately represent the typical localisation pattern of GFP-WDR48. In addition, we have\nprovided a series of confocal images of GFP-WDR48 acquired at different focal planes as\nSupplementary Fig. 8, to further demonstrate its subcellular localisation and to ensure transparency\nregarding the imaging conditions.\n3. There are several spelling errors throughout the manuscript. For instance, \"recognise\" in line 118\nshould be corrected to \"recognize,\" and \"localisation\" in line 226 should be changed to \"localization.\"\nWe thank the reviewer for pointing this out. Since Nature Communications is published by a UK-\nbased publisher, we have used British English throughout the manuscript. Therefore, spellings such as\n“recognise” and “localisation” follow British English conventions."}]

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