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. 2022 Jun 7;190(1):732–744. doi: 10.1093/plphys/kiac264

Vernalization attenuates dehydration tolerance in winter-annual Arabidopsis

Lan Chen 1,#, Pengcheng Hu 2,#, Qianqian Lu 3, Fei Zhang 4, Yanhua Su 5,†, Yong Ding 6,✉,†
PMCID: PMC9434170  PMID: 35670724

Abstract

In winter-annual plants, exposure to cold temperatures induces cold tolerance and accelerates flowering in the following spring. However, little is known about plant adaptations to dehydration stress after winter. Here, we found that dehydration tolerance is reduced in winter-annual Arabidopsis (Arabidopsis thaliana) after vernalization. Winter-annual Arabidopsis plants with functional FRIGIDA (FRI) exhibited high dehydration tolerance, with small stomatal apertures and hypersensitivity to exogenous abscisic acid. Dehydration tolerance and FLOWERING LOCUS C (FLC) transcript levels gradually decreased with prolonged cold exposure in FRI plants. FLC directly bound to the promoter of OPEN STOMATA1 (OST1) and activated OST1 expression. Loss of FLC function resulted in decreased dehydration tolerance and reduced OST1 transcript levels. FLC and OST1 act in the same dehydration stress pathway, with OST1 acting downstream of FLC. Our study provides insights into the mechanisms by which FRI modulates dehydration tolerance through the FLC–OST1 module. Our results suggest that winter-annual Arabidopsis integrates dehydration tolerance and flowering time to adapt to environmental changes from winter to spring.

Prolonged exposure to cold temperatures gradually attenuates dehydration tolerance via the FLOWERING LOCUS C–OPEN STOMATA1 module in winter-annual Arabidopsis.

Introduction

Plants are often exposed to biotic and abiotic stresses, and drought stress strongly affects plant growth and productivity (Reddy et al., 2004). Plants respond to stress conditions with a variety of biological signals at appropriate times (Takahashi and Shinozaki, 2019; Takahashi et al., 2020). For example, the phytohormone abscisic acid (ABA) plays a prominent role in abiotic stress tolerance and plant development. Ongoing work has identified components of the ABA signaling pathway. OPEN STOMATA 1 (OST1, also known as SnRK2.6 and SnRK2E), which belongs to the Sucrose nonfermenting 1-related protein kinase 2 (SnRK2) subfamily, is critical for stomata opening (Mustilli et al., 2002; Yoshida et al., 2002; Fujii et al., 2011). OST1/SnRK2.6/SRK2E is preferentially expressed in guard cells, and a mutation in OST1 results in defects in stomatal closure in response to ABA treatment (Mustilli et al., 2002). SnRK2.2/SRK2D and SnRK2.3/SRK2I are closely related to OST1; these SnRK2s interact with protein phosphatase 2Cs (PP2Cs) including ABA INSENSITIVE 1 (ABI1), ABI2, and HOMOLOG OF ABI1 (Vlad et al., 2009; Soon et al., 2012). In the absence of ABA, PP2Cs are associated with SnRK2s and inactivate SnRK2s through dephosphorylating the activation loop in the SnRK2 (Soon et al., 2012; Zhu, 2016).

ABA molecules are perceived by ABA receptors when plants experience a water deficit. Upon ABA perception, the PYRABACTIN RESISTANCE1 (PYR1)/PYR1-LIKE (PYL)/REGULATORY COMPONENTS OF ABA RECEPTORS (RCAR) family of ABA receptors change their conformation, which enables them to associate with and inhibit PP2Cs, leading to activation of SnRK2s (Fujii et al., 2009; Ma et al., 2009; Park et al., 2009; Hauser et al., 2017). There are 14 PYR/PYL/RCAR genes in Arabidopsis (Arabidopsis thaliana), and 6 of them (PYR1, PYL1, PYL2, PYL4, PYL5, and PYL8) are expressed in guard cells and are relevant for stomatal closure (Gonzalez-Guzman et al., 2012; Dittrich et al., 2019). The pyr1 pyl1 pyl2 pyl4 pyl5 pyl8 sextuple mutants display large stomata, rapid water loss, and an ABA-insensitive phenotype, similar to snrk2.2 snrk2.3 ost1 mutants (Gonzalez-Guzman et al., 2012).

SnRK2s are activated by Raf-like kinases (RAFs), and activated SnRK2s intermolecularly trans-phosphorylate other SnRK2s (Lin et al., 2020, 2021). Once activated, SnRK2s directly phosphorylate numerous target proteins involved in ABA responses and stomatal movement, including the ABA-responsive element (ABRE) binding (AREB) proteins/ABRE-binding factor (ABF) family of transcription factors such as RELATED TO ABI3/VP1 (RAV1), and ion channels such as SLOW ANION CHANNEL-ASSOCIATED 1 (SLAC1) (Zhu, 2016; Hauser et al., 2017). AREB1/ABF2, AREB2/ABF4, and ABF3 are bZIP domain proteins that directly bind to ABREs to regulate response genes (Uno et al., 2000; Fujita et al., 2013). The areb1 areb2 abf3 triple mutants exhibit increased ABA insensitivity and decreased tolerance to drought stress (Fujita et al., 2009; Yoshida et al., 2010, 2015). RAV1 binds to the promoters of ABI3, ABI4, and ABI5, and represses their transcription (Feng et al., 2014). RAV1 is phosphorylated by SnRK2s, and phosphorylated RAV1 relieves the transcriptional repression of ABI5 (Feng et al., 2014).

In Arabidopsis, flowering times vary considerably among accessions. These accessions were generally classified into summer annuals and winter annuals based on their flowering behavior (Gazzani et al., 2003). Summer annuals complete their life cycle in a single growing season with rapid flowering, whereas in winter annuals the prolonged cold in winter is necessary to accelerate flowering. Winter-annual plants require vernalization, which prevents a premature transition to the reproductive phase before the threat of freezing stress has passed (Chinnusamy et al., 2007). Winter-annual Arabidopsis accessions contain active alleles of the FRIGIDA (FRI) gene, whereas rapid-cycling accessions contain defective FRI (Michaels and Amasino, 1999; Johanson et al., 2000).

FLOWERING LOCUS C (FLC) encodes a MADS-box domain protein that acts as a repressor of flowering (Michaels and Amasino, 1999). FRI upregulates FLC transcription, while vernalization antagonizes the ability to silence FLC transcription. Genetic screening for early-flowering mutants in winter-annual backgrounds identified many genes that are required for FLC expression. Among these genes, SUPPRESSOR OFFRI4 (SUF4) encodes a nuclear-localized zinc-finger protein that specifically binds to the FLC promoter (Kim et al., 2006; Kim and Michaels, 2006). SUF4, FRI, and other FLC-specific regulators, FRIGIDA like 1 and FRI ESSENTIAL1, form a stable complex to activate the transcription of FLC (Choi et al., 2011; Li et al., 2018). Mutations in SUF4 reverse the late-flowering time of FRI and repress FLC transcription (Kim et al., 2006; Kim and Michaels, 2006).

Plants that have experienced dehydration and heat stress will exhibit a rapid and strong response to the next occurrence of these stresses, and these effects generally persist for around 1 week (Ding et al., 2012, 2013; Friedrich et al., 2021). Cold temperature affects the remodeling of cell and tissue structures and the reprogramming of metabolism, gene expression, and cytosolic calcium ion concentration. Membrane fluidity, absorption rate of water, water content of leaves, and nutrient uptake decrease under cold stress (Kuwagata et al., 2012), suggesting that cold stress might also trigger a water stress response. In addition to changes in metabolism and responsive genes, the prolonged cold in winter also functions as a signal to promote flowering in the next spring. This is known as an epigenetic memory of winter cold (Whittaker and Dean, 2017; He and Li, 2018). These results hint that cold temperatures induce short-term and long-term physiological changes. However, it is unknown whether low temperatures during winter lead to changes in stress responses in the next spring.

Here, we report that dehydration tolerance is attenuated after vernalization in winter-annual Arabidopsis but not in summer-annual Arabidopsis, and reveal how the FLC–OST1 module allows winter-annual Arabidopsis to adapt to environmental changes from winter to spring.

Results

Winter-annual Arabidopsis displayed high dehydration tolerance

To investigate dehydration tolerance, we chose two winter-annual Arabidopsis accessions containing functional FRI. Specifically, the summer-annual accession Columbia-0 (Col-0) transformed with transgenes that provide a functional FRI fused to a gene encoding green fluorescent protein (GFP) and the Col-0 genotypes crossed with FRI from the FRI-SF-2 background were termed FRIGFP and FRISF2, respectively (Michaels and Amasino, 1999; Johanson et al., 2000; Sung and Amasino, 2004; Hu et al., 2014; Lu et al., 2017). The FRIGFP and FRISF2 plants displayed a late-flowering phenotype with high induction of FLC (Supplemental Figure S1, A–C), showing that they successfully repressed flowering in Col-0, similar to a winter-annual accession.

We examined water loss in detached leaves, and the water loss rates of FRIGFP and FRISF2 plants were slower than that of wild-type (WT) Col-0 plants (Figure 1A). To induce osmotic stress, we then treated 4-day-old seedlings with mannitol for another 15 days. The leaves of FRIGFP and FRISF2 plants were mostly green whereas those of WT plants became yellow (Figure 1B). Soil-grown 14-day-old FRIGFP and FRISF2 plants were less sensitive to water withdrawal than WT plants (Figure 1C). The survival rate of FRIGFP and FRISF2 plants was over 40% after 16 days of water withdrawal and 3 days of re-watering, whereas it was only 20% for WT plants (Figure 1, C and D). FRI plants exhibited high tolerance to dehydration stress, which could be explained, at least partly, by the slow rate of water loss in FRI leaves. The slow rates of water loss suggested a possible difference in how these plants regulate stomatal closure.

Figure 1.

Figure 1

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Response of FRI plants to dehydration stress. A, Water loss in WT, FRIGFP, and FRISF2 plants. Water content was measured in detached leaves after air-drying for the indicated time and was measured as a percentage of FW. B, Representative images of FRIGFP and FRISF2 plants after mannitol treatment. Four-day-old seedlings were transformed to 1/2 MS medium containing different concentration of mannitol and grown for 15 days. C, Images of plants under different water treatments. Fourteen-day-old plants were grown with continuous water (Water) or were grown without water for 16 days (No water) and then re-watered for 3 days (Re-watered). D, Survival ratio of WT, FRIGFP, and FRISF2 plants with water and re-watering. E, Images of representative stomata of WT, FRIGFP, and FRISF2 plants grown under different concentrations of ABA. Bar = 5 µm. F, Widths of stomatal apertures from WT, FRIGFP, and FRISF2 plants grown under different concentrations of ABA. Sixty stomata were scored for each genotype. G, Relative widths of stomatal apertures from WT, FRIGFP, and FRISF2 plants expressed as the width under different concentrations of ABA relative to the width without ABA. H, Representative image of FRIGFP and FRISF2 plants with ABA treatment. Four-day-old seedlings were transformed to 1/2 MS medium containing different concentrations of ABA for 10 days. Measurements are means ± sd [n = 4 for (A) and (D), n = 3 for (F) and (G); where n is the number of independent experiments]. Asterisks indicate significant differences (*P < 0.05 and **P < 0.01) according to t tests.

FRI is involved in the ABA signaling pathway

To examine stomatal closure in the FRI leaves, we measured the stomatal apertures. The FRIGFP and FRISF2 leaves had small stomatal apertures (Figure 1, E and F). We induced stomatal closing with ABA and observed rapid closing rates in FRIGFP and FRISF2 plants (Figure 1, F and G), which indicates that FRI is involved in the ABA signaling pathway.

To test whether FRI affects the ABA signaling pathway, we treated FRI seedlings with ABA. FRIGFP and FRISF2 seedlings were mostly yellow after 14 days ABA treatment, whereas WT seedlings were green (Figure 1H). We then investigated the germination rate and cotyledon-greening percentage. In the absence of ABA, germination was similar among FRIGFP, FRISF2, and WT plants. In the presence of ABA, seed germination rates were attenuated in FRI and WT plants. However, compared with WT plants, FRIGFP and FRISF2 plants exhibited high sensitivity to ABA (Supplemental Figure S2, A and B). Accordingly, the cotyledon-greening percentages also suggested that FRIGFP and FRISF2 plants are more sensitive to ABA than WT plants (Supplemental Figure S2, C and D). Taken together, these results indicate that FRI is involved in the ABA signaling pathway.

Dehydration tolerance in FRI plants gradually decreases during vernalization

We next investigated dehydration tolerance in FRI plants during vernalization. To this end, we treated 7-day-old seedlings with vernalization (at 4°C) for 2, 4, and 8 weeks, then grew plants in soil for another 2 weeks before performing a water-loss assay. The water loss rates of detached leaves were rapid after vernalization in FRISF2 and FRIGFP plants, but not in WT plants (Figure 2A and Supplemental Figure S3, A and D). We observed rapid water loss rates in FRI plants exposed to 8 weeks vernalization, medium water loss rates in FRI plants exposed to 4 weeks vernalization, and low water loss rates in FRI plants exposed to 2 weeks vernalization (Figure 2A and Supplemental Figure S3A). These results suggest that cold exposure attenuates dehydration tolerance in FRI plants and that water loss rates in FRI plants are associated with periods of cold exposure during vernalization.

Figure 2.

Figure 2

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Water loss rates of FRISF2 plants under prolonged cold treatment. A, Water loss in FRISF2 plants under prolonged cold treatment. Water content was measured after air-drying detached leaves for the indicated time and is expressed as a percentage of FW. 0, 2, 4, and 8 weeks vernalization were indicated with 0, 2, 4, and 8 W, respectively. B, Representative stomata of FRISF2 plants grown under prolonged cold treatment. Bar = 5 µm. C, Widths of stomatal apertures from FRISF2 plants grown under prolonged cold treatment. Sixty stomata were scored for each genotype. Measurements are means ± sd [n = 3 for (A) and (C), where n is the number of independent experiments]. Asterisks indicate significant differences (*P < 0.05 and **P < 0.01) according to t tests.

We next investigated the stomatal apertures in FRI plants with and without vernalization. There were no substantial differences in the widths of stomatal apertures among FRI plants with no vernalization and those with 2, 4, and 8 weeks vernalization (Figure 2, B and C and Supplemental Figure S3, B and C). This suggests that vernalization is not involved in stomata development. We then investigated the response of FRI plants to ABA during vernalization. The stomatal widths of FRI plants were reduced in the presence of ABA. However, stomatal closure was impaired when the period of exposure to cold was extended to 2, 4, and 8 weeks (Figure 2, B and C and Supplemental Figure S3, B and C). These results suggest that prolonged exposure to cold gradually reduces sensitivity to ABA in FRI plants.

FRI promotes FLC transcription but vernalization silences FLC transcription. Reverse transcription quantitative PCR (RT-qPCR) results revealed that transcription levels of FLC were gradually reduced in FRISF2 plants but not in WT plants during vernalization (Supplemental Figure S3, E and F). The reduced transcription level of FLC was accompanied by a decrease in dehydration tolerance (Figure 2, A−C and Supplemental Figure S3D), suggesting that FLC might be involved in dehydration tolerance. FLC was highly induced in FRISF2 plants and moderately induced in FRIGFP plants (Supplemental Figure S1C). Compared with FRIGFP plants, FRISF2 plants exhibited increased dehydration tolerance (Figure 1, A–G). Plants with high transcript levels of FLC exhibited enhanced dehydration tolerance, suggesting that FLC might be positively linked to dehydration tolerance.

FLC is epistatic to FRI in dehydration tolerance

To investigate the function of FLC in dehydration tolerance, we examined the water loss of flc mutants in detached leaves. We observed rapid water loss in flc-2 and flc-3 plants (Figure 3A). We then examined stomatal apertures. The flc plants had stomatal apertures with similar widths to those of WT plants. However, the stomatal widths of flc plants were larger than those of WT plants in the presence of ABA (Figure 3, B–D), suggesting that mutations in FLC reduce stomatal closure in response to ABA. We next treated flc seedlings with mannitol. The leaves of flc-2 and flc-3 plants were mostly yellow whereas WT leaves were largely green (Figure 3E). In a soil drought experiment, the survival ratio of flc-2 and flc-3 plants was less than that of WT plants after 14 days of water withdrawal and 3 days of re-watering (Figure 3, F and G).

Figure 3.

Figure 3

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FLC is involved in dehydration tolerance. A, Water loss in WT, flc-2, and flc-3 plants. Water content was measured in detached leaves after air-drying for the indicated time and was expressed as a percentage of FW. B, Representative stomata of flc-2, and flc-3 mutants under different concentrations of ABA. Bar = 5 µm. C, Widths of stomatal apertures from WT, flc-2, and flc-3 plants under different concentrations of ABA. Sixty stomata were scored for each genotype. D, Relative widths of stomatal apertures from WT, flc-2, and flc-3 plants expressed as the width under different concentrations of ABA relative to the width without ABA. E, Representative images of seedlings treated with mannitol. Four-day-old seedlings were transformed onto 1/2 MS medium containing mannitol for 10 days. F, Representative images of plants under different water treatments. Fourteen-day-old plants were grown with continuous water (Water) or were grown without water for 16 days (No water) and then re-watered for 3 days (Re-watered). G, The survival ratio of WT, flc-2, and flc-3 plants with water and re-watering was shown. H, Water loss in WT, flc-3, and FRISF2 flc-3 plants. Water content was measured in detached leaves after air-drying for the indicated time and was expressed as a percentage of FW. Measurements are means ± sd [n = 3 for (A), (G), and (H), n = 4 (C) and (D); where n is the number of independent experiments]. Asterisks indicate significant differences (*P < 0.05 and **P < 0.01) according to t tests.

To investigate the genetic relationship between FLC and FRI in the dehydration stress pathway, we crossed flc-3 into FRISF2 and generated FRISF2 flc-3 double mutants. The water loss rates of FRISF2 flc-3 mutants were similar to those of flc-3 plants (Figure 3H), suggesting that FLC is epistatic to FRI in dehydration tolerance.

SUF4 is required for stomatal closure

SUF4 and FRI form a protein complex and promote FLC transcription; therefore, we speculated whether SUF4 was involved in dehydration tolerance. To test this, we induced osmotic stress by treating suf4-2 and suf4-4 seedlings with different concentrations of mannitol, after which suf4-2 and suf4-4 leaves were yellow (Figure 4A). Then, we induced drought stress by withdrawing water and observed rapid water loss in suf4-2 and suf4-4 plants (Figure 4B). The survival rate of WT plants was over 60%, whereas only 25% of the suf4-2 and suf4-4 plants recovered after 14 days of water withdrawal and 3 days of re-watering (Figure 4, C and D). Finally, the stomatal widths of suf4-2 and suf4-4 plants were similar to those of WT plants without ABA, and the stomatal widths of suf4-2 and suf4-4 plants were greater than those of WT plants in the presence of ABA (Figure 4, E and F). These results suggest that SUF4 is involved in stomatal closure via the ABA signaling pathway.

Figure 4.

Figure 4

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Response of suf4 to dehydration stress. A, Representative images of plants after mannitol treatment. Four-day-old seedlings were transformed to 1/2 MS medium containing mannitol for 10 days. B, Water loss in WT, suf4-2, and suf4-4 mutants. Water content was measured in detached leaves after air-drying for the indicated time and was expressed as a percentage of FW. C, Images of plants under different water treatments. Fourteen-day-old plants were grown with continuous water (Water) or were grown without water for 16 days (No water) and then re-watered for 3 days (Re-watered). D, Survival ratio of WT, suf4-2, and suf4-4 plants with water and re-watering. E, Representative images of stomata of suf4 mutants grown under different concentrations of ABA. Bar = 5 µm. F, Widths of stomatal apertures from WT, suf4-2, and suf4-4 plants grown under different concentrations of ABA. Sixty stomata were scored for each genotype. G, Water loss in suf4-2, flc-3, and suf4-2 flc-3 plants. Water content was measured in detached leaves after air-drying for the indicated time and was expressed as a percentage of FW. Measurements are means ± sd [n = 3 for (B), (D), (E), (F), and (G)]. Asterisks indicate significant differences (*P < 0.05 and **P < 0.01) according to t tests.

To investigate the relationship between SUF4 and FLC in dehydration stress, we crossed suf4-2 into flc-3 and generated suf4-2 flc-3 double mutants. The water loss rate of suf4-2 flc-3 plants was similar to that of flc-3 plants (Figure 4G), suggesting that SUF4 and FLC are involved in the same dehydration stress pathway.

FLC directly binds to the promoter of OST1 in vitro and in vivo

Given that FLC promotes stomatal closure and improves dehydration tolerance, we next examined dehydration response genes. The transcript levels of RD29A, RD29B, COR15A, RAB18, RD22, and RD26 were induced in WT plants after 7 days soil drought (Supplemental Figure S4). However, the high induction of RD29A, RD29B, and COR15A but not RAB18, RD22, and RD26, was impaired in flc mutants (Supplemental Figure S4).

Since RD29A and RD29B contain ABA-responsive elements that are activated by ABA and drought (Uno et al., 2000; Narusaka et al., 2003), we then investigated the transcript levels of genes in the ABA signaling pathway. The transcript levels of OST1, SnRK2.2, and SnRK2.3 were induced under drought conditions in WT but not in flc plants (Figure 5A). Low levels of OST1 transcript were observed in flc plants under both well-watered and dehydration stress conditions, suggesting that FLC might be essential for OST1 activation (Figure 5A). The transcript levels of SnRK2.2 and SnRK2.3 were lower in flc plants than in WT plants under dehydration, but not under well-watered conditions (Figure 5A).

Figure 5.

Figure 5

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FLC binds to the promoter of OST1. A, Transcript levels of OST1, SnRK2.2, and SnRK2.3 in WT and flc plants grown under a well-watered condition or 7 days soil drought treatment. Experiments were repeated at least three times, and data from the representative experiments are presented as means ± se (n = 3 replicates). Asterisks indicate significant differences (*P < 0.05 and P < 0.01) according to t tests, ns indicates no significance. B, Gene structure of OST1 indicating exons (boxes), introns (lines), and conserved binding motif of FLC (triangle). The primers used for ChIP-qPCR analysis are marked. C, Sequences of probes used for EMSA. The conserved sequence of CCAATAAAAAGCCC is shown in blue, and the mutated nucleotides are shown in red. D and E, Gel shift assay of FLC and various probes. The ability of FLC to bind to WT and mutated probes labeled with P32 were examined (D), and the binding specificity was tested by adding unlabeled WT competitor probe or mutated probes (E). F, Enrichment of FLC at the OST1 promoter. The ChIP-qPCR primers are indicated in (B). Experiments were repeated at least 3 times, and data from the representative experiments are presented as means ± se (n = 3 replicates). Asterisks indicate significant differences (*P < 0.05 and **P < 0.01) according to t tests.

Next, we tested whether FLC can directly bind to these genes. First, we examined the promoter sequences of these genes to look for binding motifs. The promoter of OST1 contains a conserved FLC binding motif (Hepworth et al., 2002; CCAATAAAAAGAAA) (Figure 5B and Supplemental Figure S5A). We, therefore, examined the direct binding of FLC to fragments of the OST1 promoter using an electrophoretic mobility shift assay (EMSA). We observed the FLC band with the WT probe but not with the mutated probes except mutated probe 4 (Figure 5, C and D), suggesting that FLC could directly bind to the promoter of OST1. To confirm these results, we used the WT and mutated probes as competitor probes. The EMSA showed retarded bands in the presence of mutated competitor probe 1, mutated competitor probe 2, and mutated competitor probe 3, but not when the reaction included WT competitor probes and mutated competitor probe 4 (Figure 5E). We also found two atypical FLC binding motifs in the promoter of OST1, and EMSA results revealed that FLC could directly bind with these motifs (Supplemental Figure S5, A–C).

To test whether FLC bound to the promoter of OST1 in vivo, we complemented flc-3 with FLC fused with HA driven by the 35S promoter (Pro35S:HA-FLC). The early flowering phenotype of flc-3 was rescued in transgenic plants (Supplemental Figure S5D). We measured the profiles of FLC using chromatin immunoprecipitation (ChIP) with a HA-specific antibody, followed by quantitative PCR (qPCR) analysis of the amount of DNA enrichment. FLC was highly enriched in regions 2–4 of OST1 at the promoter of OST1, where the FLC binding motifs occurred (Figure 5, B and F). These results indicate that FLC binds to the promoter of OST1in vitro and in vivo.

FLC is required for OST1 activation

We then investigated whether FLC could activate OST1 directly. The constructs containing β-glucuronidase (GUS) driven by the OST1 promoter (ProOST1:GUS) were generated and co-transformed into Arabidopsis protoplasts with or without constructs containing FLC driven by 35S (Pro35S:FLC). The ProOST1:GUS activity was highly induced in the presence of FLC (Figure 6, A and B). These results suggest that FLC can activate OST1 via its promoter. We also observed that SnRK2.2 and SnRK2.3 were slightly activated by FLC (Figure 6, A and B). Sequence analysis revealed that three atypical CArG motifs and one untypical CArG motif exist in the promoter of SnRK2.2 and SnRK2.3, respectively (Supplemental Figure S6, A and B).

Figure 6.

Figure 6

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FLC activates OST1in vitro and in vivo. A, Vectors used for GUS activity. B, GUS activity from the ProOST1:GUS, ProSnRK2.2:GUS, and ProSnRK2.3:GUS reporter constructs for cells transformed with FLC. The x-axis shows the relative GUS activity compared with the internal luciferase control (35S:LUC). Measurements are means ± sd (n = 3). C, Representative OST1-GFP signaling in ost1-3 and ost1-3 flc-3 plants with and without ABA. Bar = 50 µm. D, Relative OST1-GFP intensity in ost1-3 and ost1-3 flc-3 plants. The x-axis shows the relative GFP intensity measured by ImageJ. Fifty guard cells were scored for each background. Measurements are means ± sd (n = 3). E, Water loss in flc-3, ost1-3, and flc-3 ost1-3 plants. Water content was measured in detached leaves after air-drying for the indicated time and was expressed as a percentage of FW. Measurements are means ± sd (n = 3). F, Transcript levels of OST1 in FRISF2 plants during vernalization. RNA was isolated from seedlings after 2, 4, and 8 weeks vernalization. Experiments were repeated at least three times, and data from the representative experiments are presented as means ± se (n = 3 replicates). Asterisks indicate significant differences (*P < 0.05 and **P < 0.01) according to t tests.

To further confirm these results, we complemented ost1-3 with OST1 fused with GFP driven by the native OST1 promoter (ProOST1:OST1-GFP). The high water loss rates of ost1-3 were rescued in transgenic plants (Supplemental Figure S7A). RT-qPCR results revealed that the transcription levels of OST1 in ProOST1:OST1-GFP ost1-3 were similar to those in the WT (Supplemental Figure S7B), suggesting that ProOST1:OST1-GFP has a similar function as native OST1. We crossed ProOST1:OST1-GFP ost1-3 with flc-3 to generate ProOST1:OST1-GFP ost1-3 flc-3 plants and investigated the GFP signal with or without ABA treatment. The GFP signal was bright in guard cells in ProOST1:OST1-GFP ost1-3 plants, and this signal increased in the presence of ABA, whereas the GFP signal was dim in ProOST1:OST1-GFP ost1-3 flc-3 plants (Figure 6, C and D). These results suggest that FLC is required for OST1 activation.

To investigate the genetic relationship between FLC and OST1, we generated the flc-3 ost1-3 double mutant. The water loss rate of flc-3 ost1-3 leaves was similar to that of ost1-3, suggesting that OST1 is epistatic to FLC (Figure 6E). Since OST1 is downstream of FLC in dehydration tolerance, we speculated that one more copy of OST1 will delay the water loss rates of flc-3. To further confirm this, we crossed out ost1-3 from ProOST1:OST1-GFP ost1-3 flc-3 and generated ProOST1:OST1-GFP flc-3. The water loss rates of ProOST1:OST1-GFP flc-3 were reduced compared with flc-3 plants (Supplemental Figure S7C), suggesting that one more copy of OST1 could partially rescue the rapid water loss of flc-3. We examined the FLC expression pattern and observed GUS driven by the FLC native promoter in guard cells (Supplemental Figure S7D).

We then investigated the transcript levels of OST1, SnRK2.2, and SnRK2.3 in WT, FRI, and suf4 plants. The RT-qPCR results showed that OST1, SnRK2.2, and SnRK2.3 were induced under drought conditions in FRI and WT, but not in suf4 plants (Supplemental Figure S8, A and B). OST1 was upregulated in FRI plants and downregulated in suf4 plants under well-watered and drought conditions. SnRK2.2 and SnRK2.3 did not exhibit any difference in FRI. However, the transcript levels of SnRK2.2 and SnRK2.3 were attenuated in suf4 plants under drought conditions but not under well-watered conditions (Supplemental Figure S8, A and B).

These results are consistent with the reduction in OST1 in flc plants. Therefore, we examined the transcript level of OST1 during vernalization. The transcript levels of OST1 were slightly attenuated by 2 weeks vernalization and were substantially reduced by 4 and 8 weeks vernalization in FRI plants, but not in WT (Figure 6F and Supplemental Figure S8C). These results suggest that vernalization represses OST1 transcription. The similar patterns of FLC and OST1 transcript levels during vernalization in FRI plants suggest that FLC and OST1 work together and FLC is essential for OST1 activation.

Discussion

In this study, we demonstrated that prolonged cold conditions reduce dehydration tolerance in winter-annual Arabidopsis via the FLC–OST1 pathway. Winter-annual Arabidopsis plants containing functional FRI displayed high tolerance to dehydration, and this dehydration tolerance was gradually attenuated with prolonged cold treatment. Sensitivity to ABA was gradually attenuated in FRI plants during vernalization. Meanwhile, FLC transcript levels were highly induced in FRI plants and gradually decreased during vernalization. Loss of FLC function resulted in decreased dehydration tolerance and reduced the response to ABA. FLC directly bound to the promoter of OST1 and promoted OST1 transcription. The water loss rates of flc-3 ost1-3 double mutants were similar to those of ost1-3 single mutants, suggesting that OST1 is downstream of FLC in the dehydration stress pathway. Together, our results provided compelling evidence that prolonged exposure to cold attenuates dehydration tolerance as well as FLC and OST1 transcript levels in winter-annual Arabidopsis.

OST1 is a master regulator in the dehydration stress pathway. OST1 is inhibited by PP2Cs and activated by RAFs through phosphorylation (Fujii et al., 2009; Lin et al., 2020, 2021). In addition to AREB/ABF, RAV1, and SLAC1, OST1 also phosphorylates ICE1 and enhances its stability (Ding et al., 2015). Our study showed that vernalization reduces OST1 transcript levels and attenuates dehydration tolerance via FLC in winter-annual Arabidopsis. A reduction in FLC was associated with attenuation of OST1 during vernalization, suggesting that FLC and OST1 work together. We confirmed these results by analyzing the FLC expression pattern; we observed ProFLC-GUS in guard cells and FLC promoted OST1-GFP signals in guard cells. FLC directly bound to the promoter of OST1in vitro and in vivo and activated transcripts of OST1. FLC also bound to the promoter of FT and repressed FT transcription, suggesting that FLC has dual functions in transcription regulation. SHORT VEGETATIVE PHASE (SVP) is closely related to FLC and directly binds to the promoter of ABA catabolism genes, including CYP707A1, CYP707A3, and AtBG1 (Wang et al., 2018). Loss of SVP function attenuates drought tolerance, decreases expression of CYP707A1 and CYP707A3, and enhances expression of AtBG1 (Wang et al., 2018). The functions of FLC and SVP in transcription activation and repression might be dependent on interactions among MADS-box proteins (Li et al., 2008).

Vernalization accelerates flowering in winter-annual Arabidopsis by silencing FLC transcription (Yang et al., 2014; Whittaker and Dean, 2017). In addition to flowering acceleration, we observed attenuation of dehydration tolerance after vernalization. Although the relationship between these two physiological changes remains unclear, one possibility is that winter-annual Arabidopsis adapts to soil water conditions and cold temperatures. Water deficits generally occur in autumn (before winter) and sufficient water appears in the following spring. Arabidopsis seedlings experience a water deficit before and during winter; hence, high dehydration tolerance is required at that time. The soil water is sufficient for plant growth and flowering in spring, and high dehydration tolerance is unnecessary. Prolonged exposure to cold temperatures reduced dehydration tolerance and promoted flowering via FLC, suggesting that FLC integrated the flowering time and dehydration tolerance pathway to adapt to environmental changes. Together, our study revealed that vernalization attenuates dehydration tolerance via the FLC–OST1 module in winter-annual Arabidopsis.

Materials and methods

Plant growth conditions

The Arabidopsis (A.thaliana) ecotype Col-0 was grown at 22°C under a long-day (LD) photoperiod in a 16-h light/8-h dark cycle. For vernalization, 7-day-old seedlings were treated at 4°C with light for 2, 4, and 8 weeks (Tian et al., 2019). The mutant strains obtained from the SALK collection were as follows: suf4-2, SALK_093449; suf4-4, SALK_056285; and ost1-3, SALK_008068.

For osmotic and ABA treatment assays, 4-day-old seedlings were transferred into 1/2 Murashige and Skoog (MS) medium with different concentrations of mannitol or ABA. The seedlings were then grown at 22°C with a 12-h light photoperiod for 10 or 15 days. The experiments were repeated at least three times.

For soil dehydration assays, plants were grown at 22°C with a 12-h light photoperiod for 14 days, and were then grown under the same conditions with or without additional water for 14–16 days. Plants were then watered for 3 days and their recovery was determined by measuring color and leaf turgidity. The experiments were repeated at least three times.

Stomatal size assay

Stomatal aperture size measurements were performed as previously described (Ding et al., 2011). Briefly, stomata were opened by exposing the leaves to light and high humidity by floating the leaves on a buffer [2.5 mM 2-(N-morpholine)-ethanesulfonic acid, pH 6.15, 20 mM KCl, 1 mM CaCl2] for 2 h in a light chamber. Stomatal apertures were measured in the same buffer for another 2 h after the addition of ABA. Over 60 stomata were observed with a microscope and the widths of stomata were measured using a micro-ruler in three independent experiments. Pairwise comparisons were calculated using Student’s t tests (P < 0.05).

Water loss measurement

Water loss measurements were performed as previously described (Lin et al., 2020). Briefly, detached rosette leaves of 3-week-old plants under an LD photoperiod were placed in dishes on a bench with a light. Fresh weight (FW) was monitored at the indicated time. The experiments were repeated at least three times.

Electrophoretic mobility shift assay

The EMSA was performed as described (Su et al., 2017; Jiang et al., 2018). Briefly, 1–2 µg purified protein was mixed with 4 pmol ɣ-32P ATP-labeled probe with or without various dosages of unlabeled probe in binding buffer [10 mM Tris–HCl, pH 7.5, 100 µM KCl, 1 mM EDTA, 100 µg/mL BSA, 100 µM ZnCl2, 6% (v/v) glycerol, 1 mM DTT] for 1 h. After separation in a 4.5% native non-denaturing acrylamide gel, the gel was dried and exposed to X-ray film. The sequence of the probe is shown in Supplemental Text S1.

Promoter GUS assay and OST1-GFP assay

The constructs containing GUS driven by the OST1, SnRK2.2, and SnRK2.3 promoters were transformed into Arabidopsis protoplasts and incubation at 25°C for 12 h. The protoplasts were then transferred into lysine buffer (25 mM Tris–HCl pH 7.8, 1 mM DTT, 10% glycerol, and 1% Triton X-100). Supernatant (100 µL) was mixed with 900 µL of MUG substrates (10 mM Tris–HCl pH 8.0, containing 1 mM MUG and 2 mM MgCl2). This reaction was performed at 37°C for 30 min and stopped with 40 mM Na2CO3. GUS activity was measured using a fluorometer (HITACHI, Tokyo, Japan; U-281) with 365 nm excitation wavelength and 456 nm emission wavelength. The LUC activity was measured using the GloMax 96 Luminometer system (Promega, Madison, Wisconsin, USA; E6501) with LUC mix (Promega, E1980).

For the promoter GFP assay, leaves from 7-day-old seedlings were scanned using confocal microscopy (Zeiss, Oberkochen, Germany, LSM 980).

Quantitative PCR

RNA was extracted from 10-day-old seedlings after another 7 days soil drought treatment. RT-qPCR analysis was performed with a CFX real-time PCR instrument (Bio-Rad, Hercules, California, USA) and SYBR Green mixture (Roche, Basel, Switzerland). The relative expression of genes was quantified with the 2−△△CT calculation method using Ubiquitin 10 as the reference housekeeping gene for all qPCR analyses including ChIP assays. The gene-specific primers are shown in Supplemental Text S1.

ChIP assay

ChIP assay was performed as described previously (Su et al., 2017). Briefly, 7-day-old seedlings grown under an LD photoperiod were fixed with formaldehyde and quenched in glycine. After the nuclei were extracted, Anti-HA antibody (Roche; 11867423001, lot:13500600) or control IgG serum was added to the pre-cleared supernatants for an overnight incubation at 4°C. The immunoprecipitated sample was extracted and analyzed by qPCR with the gene-specific primers shown in Supplemental Text S1.

Accession numbers

Sequence data from this article can be found in the GenBank/EMBL data libraries under accession numbers: FRIGIDIA (AT4G00650), FLC (AT5G10140), SUF4 (AT1G30970), and OST1(AT4G33950).

Supplemental data

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

Supplemental Figure S1. Phenotypes of FRIGFP and FRISF2 A. thaliana plants.

Supplemental Figure S2. Germination rates and cotyledon-greening percentages of FRI A. thaliana plants.

Supplemental Figure S3. Osmotic stress response of FRIGFP A. thaliana plants under prolonged cold treatment.

Supplemental Figure S4. Relative transcription levels of downstream genes in A. thaliana flc mutants.

Supplemental Figure S5. FLC binding motifs in the promoter of OST1.

Supplemental Figure S6. FLC binding motifs in the promoter of snRK2.2 and SnRK2.3.

Supplemental Figure S7. Complementation of ost1-3 with ProOST1:OST1-GFP.

Supplemental Figure S8. Transcript levels in FRI suf4 and WT plants.

Supplemental Text S1. Plasmids and primers.

Supplementary Material

kiac264_Supplementary_Data
Click here for additional data file. (634.7KB, pdf)

Acknowledgments

We thank professor Pengcheng Wang from Shanghai Center for Plant Stress Biology, CAS for kindly providing ost1-1 mutants and all members in our laboratory for critical reading and discussion.

Funding

This work was supported by the National Natural Science Foundation of China (U19A2021 and 31871278 to Y.D., 32000242 to Y.S., and 32000241 to H.Z.), the Strategic Priority Research Program “Molecular Mechanisms of Plant Growth and Development” of CAS (grant no. XDB27030203), and the China Postdoctoral Science Foundation (grant no. 2019M662183).

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

Contributor Information

Lan Chen, Ministry of Education, Key Laboratory for Membraneless Organelles and Cellular Dynamics; Chinese Academy of Sciences (CAS), Center for Excellence in Molecular Plant Sciences; Biomedical Sciences and Health Laboratory of Anhui Province; School of Life Sciences, Division of Life Sciences and Medicine, University of Science and Technology of China, Anhui 230027, China.

Pengcheng Hu, Ministry of Education, Key Laboratory for Membraneless Organelles and Cellular Dynamics; Chinese Academy of Sciences (CAS), Center for Excellence in Molecular Plant Sciences; Biomedical Sciences and Health Laboratory of Anhui Province; School of Life Sciences, Division of Life Sciences and Medicine, University of Science and Technology of China, Anhui 230027, China.

Qianqian Lu, Ministry of Education, Key Laboratory for Membraneless Organelles and Cellular Dynamics; Chinese Academy of Sciences (CAS), Center for Excellence in Molecular Plant Sciences; Biomedical Sciences and Health Laboratory of Anhui Province; School of Life Sciences, Division of Life Sciences and Medicine, University of Science and Technology of China, Anhui 230027, China.

Fei Zhang, Ministry of Education, Key Laboratory for Membraneless Organelles and Cellular Dynamics; Chinese Academy of Sciences (CAS), Center for Excellence in Molecular Plant Sciences; Biomedical Sciences and Health Laboratory of Anhui Province; School of Life Sciences, Division of Life Sciences and Medicine, University of Science and Technology of China, Anhui 230027, China.

Yanhua Su, Ministry of Education, Key Laboratory for Membraneless Organelles and Cellular Dynamics; Chinese Academy of Sciences (CAS), Center for Excellence in Molecular Plant Sciences; Biomedical Sciences and Health Laboratory of Anhui Province; School of Life Sciences, Division of Life Sciences and Medicine, University of Science and Technology of China, Anhui 230027, China.

Yong Ding, Ministry of Education, Key Laboratory for Membraneless Organelles and Cellular Dynamics; Chinese Academy of Sciences (CAS), Center for Excellence in Molecular Plant Sciences; Biomedical Sciences and Health Laboratory of Anhui Province; School of Life Sciences, Division of Life Sciences and Medicine, University of Science and Technology of China, Anhui 230027, China.

 

Y.D. and L.C. conceived the study and designed the experiments. L.C. performed most of the experiments. P.H. performed the EMSA assay. Q.L., F.Z., and Y.S. prepared the materials with L.C. Y.D. wrote the manuscript.

The author responsible for distribution of materials integral to the findings presented in this article in accordance with the policy described in the Instructions for Authors (https://academic.oup.com/plphys/pages/general-instructions) is: Yong Ding (dingyong@ustc.edu.cn).

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Supplementary Materials

kiac264_Supplementary_Data
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