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. 2026 Apr 19;49(8):5421–5434. doi: 10.1111/pce.70549

The Circadian Oscillator Affects Both Glutathione Homeostasis and Its Response to Salt Stress in Arabidopsis thaliana

Desiré Cano‐Yelo 1, Raquel López‐Martínez 1, Francisca Sevilla 1, Alex A R Webb 2, Ana Jiménez 1, María Carmen Martí 1,✉
PMCID: PMC13353549  PMID: 42001398

ABSTRACT

Glutathione, including reduced glutathione (GSH) and oxidized glutathione (GSSG), is the major non‐protein thiol‐based redox buffer. There are diurnal rhythms in total glutathione level and in this work, we investigate whether these oscillations might be indicative of circadian regulation of total glutathione homeostasis. We have not only taken a look and determined transcript levels involved in glutathione homeostasis, but we have also measured GSH and GSSG levels, and glutathione reductase (GR) activity in both wild‐type plants and plants that constitutively express CIRCADIAN CLOCK ASSOCIATE1 (CCA1‐ox), under diurnal and circadian cycles, and under stress saline conditions. In spite of the presence of diurnal oscillations in several genes involved in glutathione biosynthesis and degradation and in GSH levels, GSH content does not appear to be regulated by the circadian oscillator as shown by the data in CCA1‐ox plants. Unlike GSH, we report that GSSG levels and GR gene expression and activity are affected by alterations in the circadian oscillator, demonstrating that Arabidopsis circadian clock modulates the homeostasis of glutathione via GR. Additionally, we show that GR response to saline stress is lack in CCA1‐ox plants, suggesting that the oscillator regulates glutathione response to salinity.

Keywords: circadian clock, glutathione, glutathione reductase, salt stress

Summary statement

The circadian oscillator affects glutathione homeostasis, more specifically GSH recycling, by affecting not only GR gene expression, but also GR total activity and GSSG levels under normal and saline conditions in Arabidopsis thaliana.

Abbreviations

ASC

reduced ascorbate

CCA1

CIRCADIAN CLOCK ASSOCIATE1

DHA

dehydroascorbate

GR

glutathione reductase

GSH

reduced glutathione

GSSG

oxidized glutathione

MS

mass spectrometry

qPCR

real‐time quantitative PCR

ROS

reactive oxygen species

TOC1

TIMINING OF CAB1

WT

wild‐type

1. Introduction

Because plants are sessile organisms, they cannot escape from changes of the environment, such as daily light and temperature cycles that emerge of the rotation of the Earth, and the adverse situations derived from abiotic or biotic stresses. Therefore, they have evolved intricate mechanisms that allow them to cope with these situations. Plant physiological processes and biotic or abiotic stresses, including salinity, lead to the formation of reactive oxygen species (ROS). ROS can act as second messengers in many signal transduction cascades or, if they are present in an excessive concentration can lead to oxidative damage at cellular level (Foyer and Noctor 2013; Dumanović et al. 2021; Hasanuzzaman et al. 2021; Kesawat et al. 2023). Enzymatic and non‐enzymatic antioxidant systems control ROS levels (Mhamdi and Van Breusegem 2018; Martí et al. 2020). Among them, the ascorbate‐glutathione (ASC‐GSH) cycle, which includes glutathione reductase (GR) (Jiménez et al. 1997; Foyer et al. 2005) eliminates H2O2. GSH and ascorbate (ASC) are crucial antioxidant molecules for plant defense against oxidative stress (Foyer and Kunert 2024). ASC is used as substrate for ASC peroxidase (APX) which then catalyzes the H2O2 detoxification. Glutathione is the major non‐protein thiol‐based redox buffer in most animals, plants, and prokaryotes, being also involved in many vital functions, such as transport and storage of reduced sulfur, detoxification of xenobiotics via glutathione‐S‐transferase (GST), and protection against heavy metals as a precursor in the synthesis of phytochelatins (Singh et al. 2016; Kumar and Trivedi 2018; Jobe et al. 2019). Glutathione is of critical importance for plant survival, thus Arabidopsis knockout mutants for glutathione biosynthesis gene GSH2 result in lethal phenotypes at embryo‐stage and early seedings (Marty et al. 2009). Under unstressed conditions, glutathione is present mainly in its reduced form GSH. ROS detoxification, especially during adverse or stress conditions, through the ASC‐GSH cycle leads to the formation of its oxidized form GSSG. GSSG is then reduced by GR using NADPH as co‐factor, closing the cycle and functioning to prevent the accumulation of H2O2 and the maintenance of cellular redox balance. GR is a member of the FAD‐binding disulfide reductase superfamily and the major enzyme responsible for reduction of GSSG to GSH in most organisms. The Arabidopsis thaliana genome encodes two GR genes, GR1 (AT3G24170) and GR2 (AT3G54660). GR1 is located in the cytosol, nuclei, and peroxisome, and GR2 is targeted to chloroplasts and mitochondria. GR2 is essential for plant development and GR1 is responsive to stress (Tzafrir et al. 2004; Zimmermann et al. 2004; Marty et al. 2009).

Together with antioxidant systems such as ASC‐GSH cycle, another important mechanism that helps plants to overcome stressful situations is the circadian oscillator. Circadian oscillators allow plants to anticipate and to adapt to daily light and temperature cycles by synchronizing multiple physiological and developmental responses. The oscillators might also act as a hub allowing the integration of the physiological and metabolic status of the plant (Sanchez and Kay 2016). As result, circadian oscillators allow plants to determine the optimum physiological responses to ensure plant survival and fitness. In turn, the physiological responses can provide feedback to fine tune the circadian oscillators to ensure proper regulation of the physiological and metabolic pathways (Webb et al. 2019; Buckley et al. 2023). The Arabidopsis circadian oscillator is thought to be made of transcriptional and translational feedback loops (TTFLs) comprising transcriptional feedback between dawn active Myb‐like transcription factors, a suite of PSEUDORESPONSE REGULATORS active through the day, and an evening complex of proteins that act to regulate transcription at night (Wang et al. 2022). Furthermore, epigenetic and post‐transcriptional and post‐translational modifications confer robustness to the oscillator (Seo and Mas 2014; Romanowski and Yanovsky 2015; Chen and Mas 2019). The circadian oscillator affects plant physiology and development through the regulation of gene expression. In Arabidopsis and numerous crop plants and trees species, such as rice, poplar, maize, tomato, and soybean, approximately 80% of transcripts cycle under light–dark or temperature cycles, and 30%–40% of these oscillations are circadian‐regulated in constant conditions (Harmer et al. 2000; Covington et al. 2008; Michael et al. 2008; Romanowski et al. 2020). Thus, in Arabidopsis, approximately 40%–50% of genes that are responsive to heat, osmotic stress, salinity, cold, or water deprivation are expressed rhythmically in constant light (Habte et al. 2014; Marcolino‐Gomes et al. 2014), being the circadian oscillator actively involved in the stress signaling perception and transduction pathways as recently reviewed by Xu et al. (2022). ROS and cytosolic‐free calcium [Ca2+]cyt are second messengers involved in responses to abiotic stresses. [Ca2+]cyt regulates circadian period in Arabidopsis, possible with a role in the oscillator (Martí et al. 2018; Martí et al. 2020). Similarly, ROS are also associated with circadian rhythms in plants, linking it to metabolism (Guadagno et al. 2018). Thus, recently, it has been shown that NADPH‐dependent thioredoxin reductase type C mediates the coupling of chloroplast redox rhythm with nuclear circadian clock (Paeng et al. 2025). Additionally, transcriptomic analysis has shown that dawn‐active oscillator gene CIRCADIAN CLOCK ASSOCIATED1 (CCA1) acts as a master regulator of the ROS‐responsive transcriptional network (Lai et al. 2012). The expression of genes involved in H2O2 homeostasis is under circadian regulation, most genes involved in H2O2 generation reach their maximum expression levels during the night, while the scavenging systems are more highly expressed during the day (Jiménez et al. 2021). Román et al. (2021) proposed that there are circadian rhythms of O2 .–‐regulated transcripts which are phased around dusk and have found that O2 .– acts as a metabolic signal affecting transcriptional control of the circadian oscillator in Arabidopsis. At biochemical and physiological levels, it has been reported that not only CAT gene expression, but also H2O2 levels and CAT activity are also regulated by the circadian clock (Lai et al. 2012) and that the circadian control of the transcription of VITAMIN C genes (VTCs) required for ascorbate biosynthesis, is translated into circadian rhythms of total ascorbate levels (Kiyota et al. 2006; Dowdle et al. 2007). Finally, results found in Nicotiana tabacum showed an increase in expression of the glutathione S‐transferase at the end of the light phase and it has been suggested that the expression could be under a circadian control (Sosa Alderete et al. 2018).

Despite glutathione's importance in balancing redox status of the cells, to our knowledge, there are no studies on the possible connection between the circadian oscillator and glutathione homeostasis. The reported diurnal oscillations of total glutathione levels (Schupp and Rennenberg 1988; Noctor et al. 1997; Huseby et al. 2013; Zechmann 2017), prompted us to investigate a possible connection by conducting experiments, not only at gene expression but as novelty, at biochemical levels. We report that GSH level is not regulated by the circadian oscillator. However, the circadian clock affects GSSG level and regulates GR gene expression and activity. In addition, we show that the circadian regulation of GSSG and GR participates in the glutathione homeostasis response to saline stress. All these data suggest a connection between the circadian oscillator and glutathione homeostasis and its response to salt stress.

2. Materials and Methods

2.1. Plant Material, Growth Condition, and Sampling

Arabidopsis (Arabidopsis thaliana L.) mutant lines carrying T‐DNA insertions in the GR1 gene (At3g24170), gr1‐1 (SALK_105794), and gr1‐2 (SALK_060425) were obtained from the Nottingham Arabidopsis Stock Centre (http://nasc.nott.ac.uk) and have been previously described (Marty et al. 2009; Mhamdi et al. 2010). The CCA1‐ox mutant has been previously used (Dodd et al. 2007). The background line used for the wild type and the mutants was Col‐0.

Arabidopsis seeds were surface‐sterilized with 10% (v/v) NaClO and 0.1% (v/v) Triton X‐100 for 3 min and rinsed 3 times with sterile dH2O. Surface‐sterilized seeds were plated onto 0.8% (w/v) Bactoagar plates containing ½ Murashige & Skoog (½MS) medium (pH 5.7 with 0.5 M KOH) without sucrose and stratified in the dark for 48 h at 4°C. For stress treatments of in vitro seedlings, the media was supplemented with 150 mM of NaCl and sterilized, and the seeds were sown onto the solidified media after seeds sterilization. Seeds were germinated and entrained in growth cabinets (Radiber, Spain) with a constant temperature of 19°C and 100 μmol m−2 s−1 neutral white light from LED tubes under 12 h: 12 h, light: dark (LD) cycles. For LD cycles experiments, plants were collected at the specific times of day depending of the experiment, when plants were 17‐day‐old. For 12 h: 12 h light: light (LL) cycles experiments, plants were grown for 17 days in LD and then transferred into LL conditions. After 24 h under LL, plants were collected at different time points depending on the experiment. Samples of leaves and shoots (green tissue) were rapidly frozen in liquid N2 and stored at −80°C until analysis, unless otherwise stated.

2.2. Analysis of Rhythmicity of Gene Expression and Metabolites

Gene expression data were obtained from the following datasets: LDHH‐Stitt (LDHH_ST) (Bläsing et al. 2005), LDHH‐Smith (LDHH_SM) (Smith et al. 2004), and LL12 LDHH (Harmer et al. 2000). In LDHH_ST and LDHH_SM, leaves from Col‐0 plants grown on soil under 12 h light: 12 h dark cycles during 35 and 29 days were collected, temperature was 22°C or 20°C and light intensity was 130 or 180 μmol m−2 s−1, respectively. For the circadian dataset LL12_LDHH, 7‐day‐old Col‐0 seedlings were collected. They were grown on agar supplemented with 3% sucrose at 22°C and under 12 h light: 12 h dark cycles with 60 μmol m−2 s−1 of light intensity, before being transferred to constant light (LL).

Rhythmicity analysis for both, gene expression and metabolite levels was performed using BioDare2 and the eJTK. BioDare2 is a repository for circadian, biological data, providing a platform for data sharing and period analysis (biodare2.ed.ac.uk) (Zielinski et al. 2014; Zieliński et al. 2022). Data were considered rhythmic if p was < 0.1 (*), < 0.05 (**), 0.01 (***), or 0.001 (****).

2.3. qPCR Analysis

Total RNA was extracted from green tissue of at least three biological replicates of around 20 pooled plants each, using the RNeasy Plant Mini Kit (QIAGEN). cDNA was synthesized from 500 ng RNA with the QuantiTect Reverse Transcription Kit (QIAGEN) in which, contaminating genomic DNA in RNA samples is effectively and rapidly removed with the gDNA Wipeout Buffer. The gene‐specific products were amplified using Power Green Sybr Green V PCR Master Mix 2X (Applied Biosystems) on a QuantStudio 1 Real‐Time PCR System (Applied Biosystems). Primers used are detailed in Supporting Table 1.

2.4. Enzymatic Activity

Frozen green plant tissue (0.2 g) was ground in liquid N2 and the powder was homogenized in a mortar using 50 mM potassium phosphate buffer (pH 8.0) containing 0.1 mM ethylenediaminetetraacetic acid (EDTA), 5 mM cysteine, 0.2% (v/v) Triton X‐100, 1% (w/v) soluble polyvinylpyrrolidone, and 0.1 mM phenylmethylsulphonylfluoride at 1:2 (w/v) ratio. The protein extract was centrifuged at 15 000 xg for 10 min and the supernatant was immediately filtered through a Cytiva PD SpinTrap G‐25 Desalting Columns equilibrated with the same buffer as used for homogenization. All operations were performed at 4°C. Glutathione reductase (GR; EC 1.6.4.2) activity was determined according to (Foyer and Halliwell 1976). The reaction mixture contained 25 mM sodium phosphate buffer (pH 7.8), 0.2 mM EDTA, 0.5 mM GSSG, 0.12 mM NADPH, and the enzyme extract. The NADPH oxidation was monitored by the decrease in the absorbance at 340 nm using a spectrophotometer Jasco V‐630 and the assay was performed at 30°C for 2 min.

2.5. Determination of GSH, GSSG, ASC, and DHA

The protocol to determine GSH, GSSG, ASC, and DHA was based on Rellán‐Álvarez et al. (2006). The method uses an extraction solution of very low pH to stop enzymatic activities, thereby avoiding the degradation of thiols, and also preventing GSH autooxidation. In addition, this method allows direct and simultaneous determination of reduced and oxidized glutathione and ascorbate by liquid chromatography‐electrospray/mass spectrometry. The extraction of the metabolites was performed as follows: for time point experiments, 0.2 g of fresh green plant tissue was collected and immediately homogenized at 4°C with cold extraction solution (5% m‐phosphoric acid (w/v) and 1 mM EDTA in 0.1% formic acid) using the tissue homogenizer Precellys Evolution (Bertin Technologies, France), and for time course experiments, samples were frozen in liquid N2 and extracted the following day. Samples were stored at −80°C and analyzed 24 h later using the methodology previously described in Calderón et al. (2018). Briefly, all determinations were performed using an Agilent Ion Trap XCT Plus Mass Spectrometer (Agilent Technologies, Santa Clara, CA) with an electrospray ionization source and coupled to an Agilent 1100 Series HPLC system (Agilent Technologies, Santa Clara, CA) at the Support Research Service (SAI, Murcia University). Stock solutions containing 100 μM of the metabolites were prepared in the same cold extraction solution. All the metabolites were determined in each sample.

2.6. Statistical Analysis

For qPCR experiments, each experiment was repeated at least 3 times with three or four biological replicates per treatment for each genotype. There were three technical replicates per sample on the plate. For GR activity measurements, experiments were performed 3 times with three or four biological replicates. Each sample was assayed at least twice. For time course GSH and GSSG determinations, assays were performed 3 times with three biological replicates. For time point ASC, DHA, GSH, and GSSG determinations, assays were performed twice with six biological replicates each. Unless otherwise stated, data are means ± SD of the biological replicates. SigmaPlot 15 program was used for the data analysis. For comparison between two groups, two‐tailed Student's t‐test was used and significant differences are expressed at p ≤ 0.05 (*), p ≤ 0.01 (**), and p ≤ 0.001 (***). In experiments that include two genotypes and two treatments, two‐way ANOVA followed by all pairwise multiple comparison procedures (Holm–Sidak method) was used to compare two groups. Single, double and triple asterisk indicate significance of p ≤ 0.05, p ≤ 0.01, and p ≤ 0.001, respectively, Holm–Sidak method. For time course experiments, rhythmicity was analyzed using BioDare2 as stated above in Section 2.2.

3. Results

3.1. The Circadian Oscillator Regulates Glutathione Homeostasis by Controlling GSSG Levels

To undertake the investigation of whether the oscillator affects glutathione homeostasis, first, we observed the abundance of the most important genes involved in GSH biosynthesis and degradation under diel (12 h light: 12 h dark cycles (LD)) and circadian free‐run (12 h light: 12 h light cycles (LL)). Second, we measured GSH and GSSG levels under LD and LL conditions.

In Arabidopsis, two genes regulate GSH biosynthesis: GSH1 (At4g23100) and GSH2 (At5g27380), being GSH1 the rate‐limiting enzyme (Noctor et al. 1996). Glutathione degradation mainly occurs in the cytosol by three members of the ChaC/GCG sub‐family of γ‐glutamyl cyclotransferases (GGCTs) encoded by GGCT2;1 (At5g26220), GGCT2;2 (At4g31290), and GGCT2;3 (At1g44790). Additionally, the apoplastic and vacuolar glutathione pools are degraded by γ‐glutamyl‐transferases (GGTs) encoded by a four genes family: GGT1 (At4g39640), GGT2 (At4g39650), GGT3 (At1g69820), and GGT4 (At4g29210) (Dorion et al. 2021). When we examined the rhythmicity of the expression of all those genes from the two LD datasets (LDHH_SM and LDHH_ST) (Smith et al. 2004; Bläsing et al. 2005) using BioDare2 (Figure 1A–I and Supporting Information Table S2), we found that GSH1, GGCT2;1, GGCT2;2, and GGT4 oscillate in 24 h LD cycles in both datasets. Additionally, other genes involved in GSH biosynthesis (GSH2) and degradation (GGT1) had 24 h rhythms only in one of the two LD datasets analyzed (Figure 1B,F and Supporting Information Table S2). GSH transcripts peak in the middle of the night, whereas GGCTs and GGTs tend to be more widely expressed across the 24 LD cycle (Figure 1A–I and Supporting Information Table S2). Thus, we further investigated whether the circadian clock is involved in the diel regulation of GSH biosynthesis and degradation gene expression by analyzing the expression of the cycling genes in LD using the same approach with the circadian dataset LL12_LDHH (Harmer et al. 2000). According to BioDare2 analysis (Figure 1A–I and Supporting Information Table S2), among all the genes considered rhythmic in LD, only GHS2, GGCT2;2, and GGT1 expression (Figure 1B,D,F and Supporting Information Table S2) maintained their rhythmic oscillation in LL, suggesting that their expression might be circadian regulated.

Figure 1.

Figure 1

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Diurnal and circadian changes in the expression of the genes involved in glutathione synthesis and degradation, and glutathione content. (A–I) Expression profile of GSH1, GSH2, GGCT2;1, GGCT2;2, GGCT2;3, GGT1, GGT2, GGT3, and GGT4 in Arabidopsis from the two LD datasets (LDHH_SM and LDHH_ST) and the circadian LL12_LDHH dataset. LD data are represented with dashed lines and LL with continuous lines. (J–L) Diurnal changes in total glutathione, GSH and GSSG content in Arabidopsis Col‐0 plants grown under 12 L:12 D cycles during 17 days before sampling. (M–O) Circadian changes in total glutathione, GSH and GSSG content in Arabidopsis Col‐0 plants grown under 12 L:12 D cycles during 17 days and transferred to LL before sampling. Dots indicate individual samples and black bars mean ± SE, n = 3. Double asterisk (**) indicates significance of p < 0.05 and triple asterisk (***) indicates significance of p < 0.01 after rhythmicity analysis using BioDare2. Yellow bars: day; black bars night; gray bars: subjective night.

In order to find out whether the diel and/or circadian regulation of gene expression might be translated to glutathione levels, we determined both, GSH and GSSG in LD and LL cycles. According to BioDare2, we found that like previous results obtained by Zechmann (2017), total glutathione content oscillated in LD cycles (Figure 1J). In addition to previous studies, that have determined total glutathione in LD cycles but not GSH and GSSG independently, we determined that diel oscillations in total glutathione levels and glutathione redox state are mainly due to oscillations of GSH (Figure 1K and Supporting Information Figure S1A) because its larger contribution to the total pool and also because rhythms for GSSG in LD were not robust along the 48 h time course according to BioDare2 (Figure 1L). These data suggest that the diel control of the expression of the genes involved in synthesis and degradation of glutathione exerts an effect at metabolite level, significantly affecting total glutathione and GSH levels, but probably not GSSG. When total glutathione, GSH, GSSG, and glutathione redox state were determined in LL (Figure 1M–O and Supporting Information Figure S1B), we only detected 24 h oscillations in GSH levels (Figure 1N).

To rule out that the lack of robust GSSG rhythms in LD and the loss of rhythms of total glutathione in LL could be due to difficulty in detection of low amplitude oscillations, and to corroborate that the oscillations detected in GSH levels in LL were depended on the circadian oscillator, we also determined the levels of GSH and GSSG in Col‐0 and compared to arrhythmic CCA1 OVEREXPRESSOR (CCA1‐ox) around 4 h after the subjective morning in the LL cycles (ZT 28 h) (Figure 2). ZT 28 h is 4 h after subjective dawn, around when total glutathione was higher in LD cycles (Figure 1J). GSH and total glutathione did not show differences in arrhythmic CCA1‐ox Arabidopsis mutant plants when compared to Col‐0 (Figure 2A,B). Interestingly, at ZT 28 h, GSSG levels were lower in CCA1‐ox plants compared to wild‐type (Figure 2C). The mutant plants contained 25% less GSSG than wild‐type. We suggest that because GSH levels are around 3–30 times higher than GSSG levels under normal conditions (Marty et al. 2009; Sobrino‐Plata et al. 2014; Wang et al. 2014; Fukushima et al. 2017; Calderón et al. 2018; Marty et al. 2019; Nguyen et al. 2019; Gasperl et al. 2021), the decrease in GSSG observed at ZT 28 h in CCA1‐ox when compared to wild‐type was not enough to account for a decrease in total glutathione levels (Figure 2A).

Figure 2.

Figure 2

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Content of glutathione and ascorbate. (A) Total glutathione, (B) GSH, (C) GSSG, (D) total ascorbate, (E) ASC, and (F) DHA levels determined in Arabidopsis Col‐0 (white) and CCA1‐ox (gray) plants grown in 12 h light: 12 h dark cycles and transferred to 12 h light:12 h light (LL) cycles at ZT0 when they were 17‐day‐old. Samples were collected after 28 h in LL. Dots indicate individual samples and black bars mean ± SD, n ≥ 6. Triple asterisk indicates significance of p ≤ 0.001 after two‐tailed Student's t‐test.

The observations regarding gene expression and GSH levels analysis suggest that external diurnal environmental cues may play a more important role than the circadian oscillator in regulating diel oscillations of glutathione. However, the differences found in GSSG levels in CCA1‐ox and wild‐type, but not in GSH, suggest that the circadian oscillator is involved in the homeostasis of glutathione, not at biosynthesis and degradation levels, but affecting GSSG content.

To validate our experimental approach, we performed two different sets of experiments. To check the determination method of GSH and GSSG, we determined both in the gr1‐1 mutant. Like described by Marty et al. (2009), we found that the Arabidopsis gr1‐1 mutant had about 4 times higher amount of GSSG than WT (Supporting Information Figure S2). Additionally, because the circadian control of the transcription of VTCs required for ascorbate biosynthesis is translated into circadian rhythms of total ascorbate levels (Kiyota et al. 2006; Dowdle et al. 2007), we decided to test whether the regulation of ascorbate biosynthesis by the oscillator could be detected by determining ascorbate in the arrhythmic clock mutant plants CCA1‐ox at ZT 28 h in LL, similarly to what we did for glutathione. Total ascorbate, ASC, and DHA levels were higher in the mutant CCA1‐ox (Figure 2D–F), in agreement to the previously described circadian regulation of ascorbate biosynthesis and the presence of circadian rhythms in ascorbate levels (Kiyota et al. 2006).

3.2. The Circadian Oscillator Regulates Expression of Glutathione Reductase Genes

GR is the major enzyme responsible for reduction of GSSG to GSH (Marty et al. 2009), therefore we studied whether the circadian oscillator could be affecting GSSG levels via GR. First, we studied whether the expression of the two GR isoforms encoded in the Arabidopsis thaliana genome, GR1 (AT3G24170) and GR2 (AT3G54660), was under diurnal or circadian regulation. We used BioDare2 on the two LD datasets LDHH_SM and LDHH_ST (Smith et al. 2004; Bläsing et al. 2005) and then, we validated those results using real‐time quantitative PCR (qPCR) in LD cycles time‐course experiments. BioDare2 results showed that both GR genes were expressed rhythmically in the two LD datasets, and that the expression peak of both genes occurred early in the morning (Figure 3A,D and Supporting Information Table S3). Consistent with BioDare2, we found by qPCR analysis that GR1 and GR2 are subjected to daily regulation and their transcript levels peak around 4 h after lights were switched on (Figure 3B,E). As a positive control, the expression of the clock gene CCA1 (AT2G46830) was also analyzed by qPCR (Figure 3C) checking that it followed the previously reported behavior (Smith et al. 2004 in their Figure 2B,C).

Figure 3.

Figure 3

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Expression profile of GR genes under LD and LL conditions. (A and D) Expression profile of GR1 and GR2 in Arabidopsis from the two LD datasets (LDHH_SM and LDHH_ST). (B, C, and E) Expression profile of GR1, GR2, and CCA1 obtained by q‐PCR in Arabidopsis Col‐0 plants grown under 12 L:12 D cycles during 17 days before sampling. Yellow bars: day; black bars: night. (F and I) Expression profile of GR1 and GR2 in Arabidopsis from the LL dataset (LL12_LDHH). (G, H, and J) Expression profile of GR1, GR2, and CCA1 obtained by q‐PCR in Arabidopsis Col‐0 and CCA1‐ox plants grown under 12 L:12 D cycles during 17 days and transferred to LL before sampling. Yellow bars: subjective day; gray bars: subjective night. Dots indicate individual samples and black bars mean ± SD, n = 3. Single (*), double (**), and triple (***) asterisk for white dots (Col‐0 plants) indicate significance of p < 0.1, p < 0.05, and p ≤ 0.01, respectively, after analysis of rhythmicity using BioDare2. According to BioDare2, rhythms in CCA1‐ox plants were not detected.

To examine whether the diel oscillations of GR1 and GR2 expression might be regulated by the circadian clock, we also examined their transcript levels using BioDare2 and the circadian dataset LL12_LDHH. BioDare2 results showed that only GR1 transcript levels were rhythmic in LL cycles with a peak occurring at the beginning of the subjective morning (Figure 3F and Supporting Information Table S3), suggesting that GR1 expression is clock‐controlled. Surprisingly, GR2 transcripts were not detected as rhythmic (Figure 3I and Supporting Information Table S3). Then, we validated the global transcriptome results by determining GR1 and GR2 transcript levels by real‐time quantitative qPCR in Col‐0 and in the arrhythmic clock mutant CCA1‐ox plants that were grown under LD cycles and transferred to constant light cycles (12 h light: 12 h light; LL). As in LD, the expression of GR1 and GR2 in Col‐0 plants under LL conditions is rhythmic according to BioDare, peaking at dawn, 4 h earlier than in LD (Figure 3G,J). In CCA1‐ox arrhythmic mutant plants, the expression of GR1 and GR2 became arrhythmic (Figure 3G,J). As a positive control for our experiments, the expression of the clock gene CCA1 was also analyzed. The results for CCA1 transcript abundance in Col‐0 and in the CCA1‐ox mutant were similar to those previously reported in LL (Wang and Tobin 1998) (Figure 3H). Additionally, to determine whether CCA1 directly regulates GR gene expression we searched for two circadian clock cis‐elements (CCA1 (AACAATCT and AAAAATCT) and EVENING ELEMENT (EE) (AAAATATCT)) in the GR gene promoters. We found the presence of CCA1 and EE motifs in the promoter sequence of GR2 but not in GR1 (Supporting Information Figure S3).

These observations indicate that a functional relationship likely exists between CCA1 and GR genes and that therefore, GR1 and GR2 expression is under the control of the circadian clock.

3.3. The Circadian Oscillator Regulates Glutathione Reductase Activity

Similar to the gene expression, we studied whether total GR activity was also affected in LD cycles. While GR1 and GR2 transcripts had large changes in abundance that peak early in the morning (Figure 3), total GR activity was only slightly higher at mid‐morning around 8 h after dawn (Figure 4A). Together our results suggest there is 4 h delay between GR peak transcript accumulation in the morning and GR activity peak later in the morning, which is sufficient to allow for translation. To investigate if there is circadian regulation of GR activity, we measured activity in WT and CCA1‐ox plants in LL, at the same time in which we previously determined GSH and GSSG levels, this is at 28 h after dawn and around 4 h after the peak of GR gene expression in these conditions. The results showed that the control that the oscillator exerts on GR gene expression also affected GR total activity. Thus, in total protein extracts from CCA1‐ox mutant line, GR activity was reduced by 20% when compared to WT plants (Figure 4B). To ensure that the assay was robustly reporting GR activity, we also performed the experiments using gr1‐1 mutant plants grown in LD cycles. As expected, gr1‐1 plants lacking GR1 (not GR2) activity showed significantly lower GR activity when compared to WT plants (Figure 4C) (Marty et al. 2009). Because GR2 is essential for plant development, gr2 null mutants are lethal in early embryo development (Marty et al. 2019) and could not be used.

Figure 4.

Figure 4

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Diurnal and circadian regulation of GR activity. (A) GR activity in Arabidopsis Col‐0 plants grown under 12 L:12 D cycles during 17 days. Dots indicate mean ± S.E.M., n = 3 independent experiments. Yellow bars: day; black bars: night. (B) GR activity in Col‐0 and CCA1‐ox plants grown under 12 L:12 D cycles during 17 days and transferred to 12 L:12 L cycles. Sampling was performed at 28 h in LL. (C) GR activity in Col‐0 and gr1‐1 plants grown under 12 L:12 D cycles during 17 days. Sampling was performed 4 h after dawn. Black bars indicate mean ± SD, n = 4 individual samples. Single and double asterisk indicate significance of p ≤ 0.05 and p ≤ 0.01, respectively, after two‐tailed Student's t‐test.

3.4. Glutathione Reductase Activity Affects Circadian Oscillator Gene Expression

To examine whether changes in GSSG levels due to GR activity might also affect the circadian oscillator, we examined the effect of the absence of GR1 activity in gr1‐1 and gr1‐2 mutants on CCA1 and TOC1 transcript abundance in LL. In the absence of GR1 activity, CCA1 and TOC1 transcript level oscillations were advanced in the mutants (Figure 5 and Supporting Information Figure S4). All these changes obtained by qPCR suggest that the period of the clock could be shorter and the clock could be slightly faster in the two mutants when compared to WT.

Figure 5.

Figure 5

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Expression profile of the clock genes CCA1 and TOC1 under LL conditions. Expression profile of CCA1 and TOC1 obtained by q‐PCR in Arabidopsis Col‐0 and gr1‐1 plants grown under 12 L:12 D cycles during 17 days and transferred to LL before sampling at 48–60 h (A) and 72–84 h (B). Dots indicate mean ± SD, n = 3. Single, double, and triple asterisk indicate significance of p ≤ 0.05, p ≤ 0.01, and p ≤ 0.001, respectively, after two‐tailed Student's t‐test. Yellow bars: subjective day; gray bars: subjective night.

3.5. The Circadian Oscillator Affects the Response of Glutathione Reductase and GSSG to Salinity

Because we have demonstrated the circadian oscillator exerts control on GR activity and GSSG levels, we tested the hypothesis that this control modulates responses to oxidative situations such as the one imposed by salt stress. We investigated the response to 150 mM NaCl. GR1 and GR2 transcript levels were slightly increased and decreased, respectively, under saline conditions in Col‐0 plants, whereas in CCA1‐ox mutant plants, salinity did not affect GR1 transcript levels (Figure 6A,B). When we determined total GR activity, we found that it increased in WT plants treated with salt, but interestingly, it was not affected in CCA1‐ox mutant plants under the same conditions (Figure 6C). One hundred fifty millimolar of salt decreased total glutathione and GSH in both WT and CCA1‐ox but salt only affected GSSG in WT plants and not in the CCA1‐ox mutant plants (Figure 6D–F). In summary, unlike in Col‐0, GR1 transcript levels, total GR activity, and GSSG were insensitive to salt treatment in CCA1‐ox, suggesting that the circadian oscillator might mediate in the response of GR to salinity. Data from Figure 6G–I indicate that this mediation is not affecting ASC and DHA levels, because 150 mM of NaCl did not affect differently ASC and DHA in Col‐0 and CCA1‐ox plants (Figure 6G–I).

Figure 6.

Figure 6

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GR expression, total GR activity and glutathione content in plants grown under control and in the presence of 150 mM NaCl. (A) Expression of GR1 and (B) GR2, (C) total GR activity and (D) total glutathione, (E) GSH, (F) GSSG, (G) total ascorbate, (H) ASC, and (I) DHA content in Arabidopsis Col‐0 and CCA1‐ox plants grown under 12 L:12 D cycles during 17 days and transferred to LL before sampling at ZT24 for expression and ZT28 for activity and metabolites determination experiments. White and gray dots represent Col‐0 and CCA1‐ox plants, respectively. Dots indicate individual samples. For (A–C), bars indicate Supporting mean ± SD, n = 3–4. For (D–I), bars indicate mean ± SD, n = 6. Single, double, and triple asterisk indicate significance of p ≤ 0.05, p ≤ 0.01, and p ≤ 0.001, respectively, after two‐way ANOVA followed by all pairwise multiple comparison procedures (Holm–Sidak method). ns: non‐significant.

4. Discussion

4.1. Diurnal Regulation of GSH and GSSG

Our data related to total glutathione and also to GSH content in Arabidopsis measured under 12 h photoperiod by HPLC‐MS (Figure 1J,K) showed a diurnal dynamic consisting of a rapid increase of total glutathione and GSH after the onset of light that peaks around 7 h after dawn, followed by a strong drop that leads to low levels of total glutathione and GSH during the night. Similarly in Arabidopsis, but under 8 h photoperiod, total glutathione content showed the same dynamic but advanced (Zechmann 2017). The same diurnal dynamic of total glutathione has been also determined in several plant species (Schupp and Rennenberg 1988; Noctor et al. 1997; Huseby et al. 2013). Additionally in Arabidopsis, immunohistochemistry and quantitative transmission electron microscopy have shown that the diurnal rhythms in total glutathione content of leaves arise from several cellular compartments that include, chloroplasts, mitochondria, peroxisomes, nuclei, and cytosol (Zechmann 2017). It has been suggested that in the dark, the lack of photorespiration leads to a shortage of the glutathione precursors glycine and cysteine. With the onset of photosynthesis, the supply of the precursors is restored and glutathione synthesis increases. The drop‐in glutathione levels has been suggested to be probably due to the increased glutathione degradation, usage for phytochelatin synthesis (Zechmann 2017), or the increased activity of GSTs (Sosa Alderete et al. 2018; Gallé et al. 2018). Our data regarding GSH content and GSH gene expression (Figure 1J–L and Supporting Information Table S2) strongly support this hypothesis. Thus, transcripts of GSH biosynthesis genes peak at similar times during the night in preparation for the GSH supply that is needed during the day. GSH content peaked at 7 h after dawn and the genes involved in the main pathways of GSH degradation peaked after the maximum levels of GSH, during a broader window of time across the 24 h LD cycle when compared with the GSH genes, probably due to the plethora of pathways that use and consume GSH.

To our knowledge, data regarding GSSG level during daily cycles have not been reported. In our study, we have shown that in the same samples where rhythms in GSH content were detected, GSSG content was lower than GSH as expected (Marty et al. 2009; Foyer and Noctor 2011; Sobrino‐Plata et al. 2014; Wang et al. 2014; Fukushima et al. 2017; Calderón et al. 2018; Marty et al. 2019; Nguyen et al. 2019; Gasperl et al. 2021), and did not display a robust and statistically significant oscillatory dynamic when BioDare2 was used as a tool to detect rhythmicity (Figure 1L), suggesting that the recycling of GSH by GR in the ASC‐GSH cycle is enough to maintain GSSG levels. Alternatively and/or simultaneously, GSH might be used in other pathways that are not accompanied by net oxidation of GSH which produces GSSG, such as the detoxification of methylglyoxal, an unavoidable toxic by‐product of glycolysis, or the post‐translational modification S‐glutathionylation (Dorion et al. 2021).

4.2. Circadian Regulation of GSH and GSSG

Our study revealed that for the main genes involved in glutathione biosynthesis and degradation, the rhythmicity that was present in LD was lost in LL for all except GSH2, GGCT2;2, and GGT1. Although according to BioDare2, we found rhythms in GSH levels in LL, we did not find rhythms in total glutathione under the same conditions (Figure 1M,N) or differences in GSH and total glutathione levels when Col‐0 was compared to CCA1‐ox (Figure 2), suggesting that the rhythmicity of GSH2, GGCT2;2, and GGT1 expression in LL might not be sufficient to support circadian cycles of glutathione and GSH and therefore, the circadian clock does not regulate glutathione biosynthesis and degradation and that GSH levels are largely driven by external environmental cues. Interestingly, we observed that GSSG levels were lower in CCA1‐ox than in Col‐0 plants in LL (Figure 2) suggesting a role of the clock controlling GSSG levels and therefore glutathione homeostasis.

On the contrary to total glutathione, we found that the levels of total ascorbate, ASC, and DHA were altered in CCA1‐ox plants (Figure 2), which agrees with the proposed role of the circadian clock regulating ASC synthesis through GDP‐l‐galactose phosphorylase (GGP) the main rate‐controlling enzyme in ASC biosynthesis (Smirnoff and Wheeler 2024). In Arabidopsis, there are two genes encoding for GGP, VTC2/AtGGP1 and VTC5/AtGGP2, being responsible for the synthesis of 80% and 20% of normal leaf ASC, respectively (Kiyota et al. 2006; Dowdle et al. 2007). VTC2 expression has a predawn phase which is altered in CCA1‐ox and cca1‐1/lhy‐11 mutants (Lai et al. 2012). Similarly, here we reported that ASC and DHA levels are altered in CCA1‐ox.

These results suggest that rhythmic oscillations in CCA1 level might be necessary for maintaining the levels of ascorbate and GSSG, but not GSH, and that statistically significant 24 h oscillations in GSSG content are difficult to detect in both LD and LL. Therefore, our data support a role of the circadian clock controlling glutathione homeostasis, not through GSH biosynthesis and degradation, but possibly as a consequence of changes in GSH oxidation or GSSG reduction rates. GR is the main enzyme responsible for GSSG reduction in plants, which prompted us to test whether GR could be a target of the circadian regulation.

4.3. Links Between GR Transcript Abundance and Activity, and the Circadian Oscillator

Our qPCR data together with the microarray data analyzed with BioDare2 showed that the two GR genes, GR1 and GR2 reach their maximum of expression after dusk in LD (Figure 3). This timing of GR transcript abundance is consistent with a role that anticipates diurnal H2O2 production that peaks around 7 h after dawn (Lai et al. 2012). The rhythmic temporal expression for both GR genes under LL in WT plants and arrhythmic expression in the CCA1‐ox mutant demonstrates GR gene expression is regulated by the circadian oscillator (Figure 3). Promoter analysis identified multiple circadian clock cis‐regulatory elements (CCA1 and EE) within the 2500‐bp of GR2 promoter, whereas no such motifs were found in the promoter region of GR1, pointing to a direct regulation of GR2 and to an indirect regulation of GR1 (Supporting Information Figure S3).

Oscillations in GR activity during the LD cycle, peaked at midday (Figure 4), the same as total glutathione and GSH in Col‐0 plants (Figure 1J,K) and similar to that of H2O2 (Lai et al. 2012). Additionally, the maximum expression of GR occurs 4 h after dusk, which allows enough time for translation (Figure 3). In the circadian arrhythmic CCA1‐ox mutant, GR gene expression became arrhythmic and GR activity was reduced (Figures 3 and 4), suggesting that the oscillator regulates both GR gene expression and GR activity. Interestingly in CCA1‐ox mutant, we have found that both GR activity and its substrate GSSG are lower than in the wild type. Whilst, this must at first seem surprising, a similar relationship has been found in CCA1‐ox catalase activity and its substrate, H2O2 (Lai et al. 2012). It might be that despite the lower GR activity in CCA1‐ox mutant, GSSG levels are lower due to the lower accumulation of H2O2 reported for CCA1‐ox mutant (Lai et al. 2012) and/or the higher levels of ASC reported here, which could be decreasing the rate of GSH oxidation through enzymatic and nonenzymatic processes.

As there is currently no experimental evidence for post‐translational regulation of GR activity (Dorion et al. 2021), it might be that the rhythmic transcriptional regulation of GR is responsible for the rhythms in activity. However, we cannot rule out the possibility that PTMs could also contribute to the rhythms of GR activity together with circadian regulation of expression. Collectively, the results suggest that the circadian oscillator mediates in glutathione homeostasis under regular growth conditions by affecting, not GSH biosynthesis and degradation, but GR gene expression and activity probably to control the concentration of GSSG for fine regulation of metabolism and stress responses.

We obtained preliminary data showing that GR activity might affect the circadian oscillator (Figure 5). Like ASC biosynthesis, that has been shown to be circadian regulated and able to affect clock gene expression (Kiyota et al. 2006; Kotchoni et al. 2009), our data suggest that GR1 activity might be also fine tuning the clock. Thus, all these data together support the idea that the circadian system not only affects ROS generation and scavenging, but also that ROS levels contribute to the fine tuning of the oscillator via ascorbate levels and maybe GR1 activity.

4.4. Link Between Glutathione Homeostasis and the Circadian Oscillator Under Saline Conditions

Soybean plants, in response to As, display a regulatory interaction between clock genes and GSH levels (Vezza et al. 2020). It has been reported that GR genes in wild‐type plants are not affected by abiotic stresses, except for GR1 that appears slightly up‐regulated in response to cold, osmotic, salt, and ultraviolet (UV) stresses in shoots (Belin et al. 2015). Whereas GR activity increased under saline conditions in Arabidopsis (Sánchez‐Guerrero et al. 2019), similar to GSSG (Sánchez‐Guerrero et al. 2021). For CCA1‐ox plants, it has been previously shown that they are hyposensitive to 5 μM methyl viologen (MV) (Lai et al. 2012). In our conditions, salt affected GR gene expression, activity and GSSG levels in WT plants but not in CCA1‐ox plants (Figure 6). The lack of response of CCA1‐ox plants suggests that a functional circadian oscillator is required for the proper GR‐mediated response of Arabidopsis to this stress situation (Figure 7). These results agree with the reduced response that CCA1‐ox plants have under MV stress and the proposed role of CCA1 in coordinating the response to oxidative stress and the regulation of ROS production and scavenging (Lai et al. 2012).

Figure 7.

Figure 7

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An illustration showing the connection between the circadian clock and glutathione homeostasis under normal and stress saline conditions.

Considering the importance of the circadian system and the glutathione function in plants, in this work, to our knowledge, for the first time, we have demonstrated that the circadian oscillator exerts its control on glutathione homeostasis, more specifically on GSH recycling, affecting not only GR gene expression, but also GR total activity and GSSG levels under normal and saline conditions. Overall, these results provide insight in how plants cope with stress, reinforcing the role of the oscillator on plant stress responses at both transcriptional and biochemical level.

Conflicts of Interest

The authors declare no conflicts of interest.

Supporting information

Supporting File 1

PCE-49-5421-s001.docx (1.2MB, docx)

Supporting File 2

PCE-49-5421-s002.docx (15.9KB, docx)

Acknowledgements

The authors want to thank Ms. Sandra Correa for the technical help. This study was supported by the following Spanish grants and fellowship: Spanish Ministry of Science, Innovation and Universities (PID2020‐119989GA‐I00/AEI/10.13039/501100011033 and FPI fellowship awarded to D.C.‐Y. (PRE2021‐099183)), Ramón Areces Foundation, i‐LINK+ International Collaboration Program (CSIC, Spain), and Séneca Foundation, Región de Murcia, Spain (22051/PI/22).

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