Next Article in Journal
Is There Daily Growth Hysteresis versus Vapor Pressure Deficit in Cherry Fruit?
Previous Article in Journal
Shoot Production and Mineral Nutrients of Five Microgreens as Affected by Hydroponic Substrate Type and Post-Emergent Fertilization
 
 
Search for Articles:
Title / Keyword
Author / Affiliation / Email
Journal
Article Type
 
 
Advanced Search
 
Section
Special Issue
Volume
Issue
Number
Page
 
Journals
Horticulturae
Volume 7
Issue 6
10.3390/horticulturae7060130
horticulturae-logo
Submit to this Journal Review for this Journal Propose a Special Issue
► Article Menu

Article Menu

share Share announcement Help format_quote Cite question_answer Discuss in SciProfiles thumb_up
...
Endorse textsms
...
Comment
first_page
settings
Order Article Reprints
Font Type:
Arial Georgia Verdana
Font Size:
Aa Aa Aa
Line Spacing:
Column Width:
Background:
Article

Role of Glutathione-Ascorbate Cycle and Photosynthetic Electronic Transfer in Alternative Oxidase-Manipulated Waterlogging Tolerance in Watermelon Seedlings

by
Jiawen Zheng
1,†,
Chunying Fang
1,†,
Lei Ru
1,
Nan Sun
1,
Yuanyuan Liu
1,
Yunping Huang
2,
Yuhong Wang
2,
Zhujun Zhu
1,* and
Yong He
1,*
1
School of Horticultural Science, Zhejiang A&F University, Hangzhou 311300, China
2
Ningbo Academy of Agricultural Sciences, Ningbo 315040, China
*
Authors to whom correspondence should be addressed.
†
J.Z. and C.F. contributed equally.
Horticulturae 2021, 7(6), 130; https://doi.org/10.3390/horticulturae7060130
Submission received: 3 May 2021 / Revised: 29 May 2021 / Accepted: 31 May 2021 / Published: 3 June 2021
Browse Figures
Versions Notes

Abstract

:
Alternative oxidase (AOX) has been documented to mitigate the oxidative stress caused by abiotic stresses. However, it remains unknown how AOX regulates the antioxidant system and photosynthesis under waterlogging. To address this issue, we used two watermelon (Citrullus lanatus L.) cultivars (waterlogging tolerant cultivar ‘YL’ and sensitive cultivar ‘Zaojia8424’) as materials and the AOX inhibitor salicylhydroxamic acid (SHAM) to investigate the effects of AOX on photosynthesis and reactive oxygen species metabolism under waterlogging. We found that waterlogging decreased leaf photosynthesis and quantum yield of photosynthesis in watermelon, and the waterlogging tolerant cultivar ‘YL’ showed higher expression level of ClaAOX than the sensitive cultivar ‘Zaojia8424’. Net photosynthesis rate was higher in ‘YL’ than ‘Zaojia8424’. Moreover, waterlogging induced photoinhibition in ‘Zaojia8424’ but not in ‘YL’. Meanwhile, waterlogging promoted the accumulation of superoxide and peroxide hydrogen, and triggered oxidative damage. ‘YL’ suffered from less severe oxidative damage due to increased contents of ascorbate, a higher ratio of reduced glutathione (GSH) to oxidized glutathione (GSSG), a higher activity of ascorbate peroxidase (APX) and catalase (CAT), and enhanced levels of CAT and APX expression, relative to ‘Zaojia8424’. However, the alleviation of photosynthesis and oxidative damage, increased content of ascorbate and higher GSH/GSSG ratio were abolished by SHAM. Our results suggested that photosynthetic electronic transfer and glutathione-ascorbate cycle are involved in waterlogging tolerance mediated by the AOX pathway in watermelon.

1. Introduction

Waterlogging is an emerging environmental factor restricting plant productivity due to climate change and influences almost 16% of global cultivated areas [1]. It is estimated that two-thirds of the total crop yield reduction worldwide has been ascribed to waterlogging during 2006–2016 [2]. Waterlogging initiates O2 deprivation within the roots, and triggers molecular, biochemical and physiological changes in both shoots and roots [1,3,4,5,6]. Leaf photosynthesis is susceptible to waterlogging. The reduction of net photosynthesis rate (Pn) by waterlogging has been observed in many plants, including cucumber (Cucumis sativus) [7], maize (Zea mays) [8], peanut (Arachis hypogaea) [9], sorghum (Sorghum bicolor) [10] and tomato (Solanum lycopersicum) [11]. Moreover, both Photosystem II (PSII) and Photosystem I (PSI) are impaired by waterlogging, represented by a decrease in the maximal photochemical efficiency of PSII and PSI [12]. Furthermore, the photosynthetic electron transport chain was interrupted by waterlogging, indicated by the reduction of effective quantum yield of PSII (YII) [13].
The chloroplast is a primary generating site of reactive oxygen species (ROS), including singlet oxygen (1O2), superoxide (O2−), hydrogen peroxide (H2O2), and the hydroxyl radical (·OH) [14,15]. The over-reduction of the photosynthetic system due to environmental stress such as waterlogging causes electron leakage from plastoquinone QA and QB to O2, and consequently results in the production of ROS [16]. It has been reported that O2− and H2O2 can be excessively produced by waterlogging in plant leaves [8,17,18], and the over-accumulation of ROS could lead to oxidative damage to lipids, proteins and nucleic acids [14].
To keep the ROS concentrations under tight control, plants have evolved a sophisticated antioxidant system, including non-enzymatic and enzymatic ROS scavenging mechanisms [19]. Enzymatic ROS scavenging mechanisms comprise superoxide dismutase (SOD), catalase (CAT) and guaiacol peroxidase (GPOD). SODs dismutate O2− into H2O2, which can be detoxified into H2O by CAT, GPOD and the glutathione-ascorbate cycle. Ascorbate and glutathione scavenge H2O2 via ascorbate peroxidase (APX) and glutathione peroxidase (GPX) [20,21]. The antioxidant system responds rapidly to waterlogging. For instance, leaf ascorbate and glutathione pools were reduced after only 3 h of anoxia in barley (Hordeum vulgare) [18].
Alternative oxidase (AOX), located in the inner membrane of mitochondria, serves as a terminal electron acceptor besides classical cytochrome c oxidase. AOX can lower ROS production in plant cells and has a positive effect on plant performance under abiotic stresses [22,23]. For instance, overexpression of AOX reduced the accumulation of O2− and prevented the oxidative damage caused by hypoxia in tobacco (Nicotiana tabacum) [24]. However, whether the glutathione-ascorbate cycle is involved in the alleviation of oxidative damage mediated by AOX under waterlogging remains unknown. Moreover, it has been reported that AOX is essential in maintaining of photosynthesis under drought stress [25,26]. However, the role of AOX in photosynthesis under waterlogging remains unclear.
Watermelon (Citrullus lanatus) is an important crop worldwide and is often threatened by waterlogging. Recently, we discovered a waterlogging tolerant cultivar of watermelon, ‘YL’, and found its tolerance to waterlogging was associated with enhanced activity of AOX via maintaining the root respiratory [27].
In this study, we examined the role of the glutathione-ascorbate cycle and photosynthesis in AOX mediated waterlogging tolerance by using the tolerant cultivar ‘YL’ and a sensitive cultivar ‘Zaojia8424’. Salicylhydroxamic acid (SHAM), an AOX inhibitor, was also employed to manipulate AOX activity. Biochemical and physiological parameters associated with the glutathione-ascorbate cycle and photosynthesis were examined.

2. Materials and Methods

2.1. Plant Genotypes and Waterlogging Treatment

Two cultivars of watermelon (Citrullus lanatus), the waterlogging tolerant cultivar ‘YL’ and a waterlogging sensitive cultivar ‘Zaojia8424’, were used in the present study. Seeds of both cultivars were surface sterilized and germinated for 3 days at 30 °C in darkness. Seedlings were transferred into 1 L plastic pots containing commercial substrate (Hangzhou Jinhai Agricultural Technology Co. LTD, China) with one seedling in each pot. All pots were placed in a growth chamber with day/night temperatures of 28 °C/20 °C, light intensity of 600 μmol m−2 s−1 and 12 h photoperiod.
After reaching the four-leaf stage, the pots were transferred to a tank filled with water to impose waterlogging treatment for 9 days. In the AOX inhibition treatment, 3 mM SHAM was sprayed on leaves every 3 days during waterlogging treatment. The second and third fully expanded leaves were harvested from the start to the end of waterlogging at an interval of 3 d (0, 3, 6 and 9 d after waterlogging stress (DAS)) to measure the contents of glutathione and ascorbate acid, and the activity of antioxidant enzymes as well. These leaves were sampled from a subset of plants, and an individual plant was sampled once. The oxidative damage and contents of superoxide and hydrogen peroxide were investigated at 9 DAS. The harvested tissues were put into the liquid nitrogen immediately and then stored at −80 °C until analyses. Twenty plants were included in each treatment and in the controls. The experiment was repeated 3 times from May to November 2020.
The shoots were harvested and weighed at the end of waterlogging.

2.2. Measurement of Photosynthesis and Chlorophyll Fluorescence

Leaf photosynthesis was measured in four plants randomly selected from each treatment with an open gas exchange system (Li-6800, Li-Cor, USA) at 9 DAS. Net photosynthetic rate (Pn), stomatal conductance (Gs), intercellular CO2 concentration (Ci) and transpiration rate (E) were determined between 9:00 and 12:00 at 800 μmol m−2 s−1 photosynthetic photon flux density. During measurement, relative humidity was maintained at 70% and leaf temperature was set at 28 ± 0.5 °C in the leaf chamber.
Chlorophyll fluorescence was measured on the same leaf for photosynthesis measurements using a pulse amplitude modulated chlorophyll fluorometer (PAM2500, Walz, Germany) at 9 DAS. Prior to Fv/Fm measurements, the leaf area assayed was dark-adapted for 20 min. The initial fluorescence (Fo) was obtained by application of a low-intensity red measuring light source, and Fm was measured after applying a saturating light pulse of 8000 μmol m−2 s−1. Minimum (F’o) and maximum (F’m) values of fluorescence in the light-adapted state at 800 μmol m−2 s−1 were also obtained in this manner. The following parameters were collected from the machine: Fv/Fm (maximum quantum efficiency of PSII photochemistry), F’v/F’m (effective quantum use efficiency of PSII in the light-adapted state), qP (proportion of open PSII), YII (the effective quantum yield of PSII), YNPQ (the quantum yield of regulated non-photochemical energy loss in PSII) and YNO (the quantum yield of non-regulated energy loss in PSII).

2.3. Assay of Electrolyte Leakage and Lipid Peroxidation in Leaves

Electrolyte leakage in leaves was assessed by an electrical conductivity meter (DDS-307, Shanghai INESA Scientific Instrument Co., Ltd., Shanghai, China) following the method of Dionisio-Sese and Tobita (1998) [28].
Lipid peroxidation was estimated in terms of thiobarbituric acid reactive substances (TBARS) according to the method of Cakmak and Marschner (1992) [29]. Briefly, 0.3 g leaves were homogenized in 2 mL 0.1% (v/v) trichloroacetic acid solution on ice. After centrifugation at 10,000× g for 5 min, the supernatant was collected and measured spectrophotometrically at 532 and 600 nm. After subtracting the non-specific absorbance at 600 nm, the TBARS concentration was determined by its extinction coefficient of 155 mM−1 cm−1.

2.4. Quantification of O2− and H2O2

For the extraction of O2−, 0.5 g watermelon leaves were ground in 2 mL potassium phosphate buffer (pH 7.8, 65 mM), and the homogenates were centrifuged at 10,000× g for 15 min at 4 °C. Then, 0.5 mL supernatants were mixed with 1.5 mL of reaction buffer (containing 3 mM hydroxylamine hydrochloride, 20 mM sulfanilamide and 2 mM α-naphthylamine in 4 M acetic acid) and incubated for 20 min at 25 °C. The absorbance of the reaction mixture was recorded at 530 nm [30].
The H2O2 content was assayed according to the method of Cheeseman (2006) [31]. Leaf tissue (0.5 g) was homogenized in 2 mL HClO4 (1.0 M) with polyvinylpyrrolidone. The homogenates were centrifuged at 10,000× g for 10 min at 4 °C. Then, 1 mL supernatant was added to 3 mL eFOX reagents, incubated for 20 min, and then the difference in absorbance between 550 and 800 nm was recorded. The H2O2 content was calculated through a standard curve.

2.5. Determination of Antioxidant Enzyme Activity

The extraction and activity determination of antioxidant enzymes were carried out as described by He et al. (2009) [32]. A 0.3 g leaf was ground with 2 mL ice-cold potassium phosphate buffer (pH 7.8, 50 mM) containing 0.2 mM EDTA and 2% polyvinylpyrrolidone, and the homogenates were centrifuged at 4 °C for 20 min at 12,000× g. Then the supernatants obtained were used for enzyme analysis.
The activity of superoxide dismutase (SOD, EC; 1.15.1.1) was assayed by its ability to inhibit the photochemical reduction of nitro blue tetrazolium [33]. The activity of catalase (CAT, EC 1.11.1.6) was quantified by monitoring the reduction of absorbance at 240 nm as the disintegration of H2O2 using the extinction coefficient 39.4 mM−1 cm−1 [29]. The activity of guaiacol peroxidase (GPOD, EC 1.11.1.7) was calculated by the increase of absorbance at 470 nm using the extinction coefficient 26.6 mM−1 cm−1 [34]. The activity of ascorbate peroxidase (APX, EC 1.11.1.11) was measured by the decrease in absorbance at 290 nm with extinction coefficient 2.8 mM−1 cm−1 [35]. Dehydroascorbate reductase (DHAR, EC; 1.8.5.1) activity was measured by the increase in absorbance at 265 nm using the extinction coefficient 14 mM−1 cm−1 [35]. Glutathione reductase (GR, EC; 1.6.4.2) activity was calculated by the decrease in absorbance at 340 nm due to NADPH oxidation using the extinction coefficient 6.2 mM−1 cm−1 [36].

2.6. Analysis of Glutathione and Ascorbate

The extraction of glutathione and ascorbate were carried out as described by Hodges et al. (1996) [37]. Briefly, 0.3 g watermelon leaf was ground in 3 mL of 5% trichloroacetic acid and the homogenate was centrifuged at 4 °C for 10 min at 12,000× g. This supernatant then was used for reduced glutathione (GSH), oxidized glutathione (GSSG) and ascorbate measurement.
For the measurement of glutathione, 0.5 mL supernatant was added to the 2 mL reaction solution containing 50 mM potassium phosphate, 0.2 mM 5,5′-dithiobis-(2-nitrobenzoic acid), 0.2 mM NADPH, and 3 units of glutathione reductase [38]. The total glutathione content was calculated by monitoring the absorbance at 412 nm for 1 min. To measure the content of GSSG, 4 μL 2-vinylpyridine was added to 200 μL extract and then the same protocol for total glutathione was used. The content of GSH was attained by deducting GSSG from total glutathione.
For the measurement of ascorbate, 0.2 mL supernatant was added to the 2 mL reaction solution containing 4% trichloroacetic acid (v/v), 22% o-phosphoric acid (v/v), 5 mM α,α’-dipyridyl and 10 mM FeCl3 and incubated for 30 min. The absorbance at 520 nm was recorded.

2.7. Identification of AOX Amino Acid Sequences from Watermelon Genome

The conserved amino acid sequence of DBD (Pfam: PF01786 for AOX) was used as a BLAST query against the watermelon genome database (http://cucurbitgenomics.org/organism/21, 1 November 2019), and the full-length amino acid sequences of the AOX proteins from arabidopsis (Arabidopsis thaliana) were obtained from National Center for Biotechnology Information (NCBI). The acquired protein sequences were confirmed by Pfam (http://pfam.xfam.org/search, 1 March 2021) and then phylogenetic analyzed with MEGA 5 and ClustalW software.

2.8. RNA Isolation and Real-Time Quantitative PCR

The second fully expanded leaves were harvested at 0, 24, 48 and 72 h after stress (HAS) to measure the expression of genes involved in antioxidant responses. Total RNA was extracted using Trizol reagent (Invitrogen, Carlsbad, CA 92008, USA) following the manufacturer’s protocol. Then, the total RNA was reverse transcribed using the THUNDERBIRD SYBR® qPCR Mix (Toyobo, Osaka, Japan). Quantitative real-time PCR was performed using SYBR Green PCR Master Mix in real-time PCR System (qTower3, Analytik, Jena, Germany). Each reaction (20 μL) contained 4 μL SYBR qPCR Mix, 0.4 μL ROX reference dye, 3 μL cDNA, 0.8 μL each of forward and reverse primers and 11 μL distilled water. PCR was run for 35 cycles of 20 s at 95 °C, 30 s at 60 °C and 30 s at 72 °C. UBCP was used as reference gene. The specific primers employed for AOX, CSD, CAT, APX, GPOD, DHAR, GR, RBoh and UBCP can be found in Table S1. The relative gene expression was calculated according to Livak and Schmittgen (2001) [39] and was normalized to the results from the expression of ‘Zaojia8424’ at 0 HAS.

2.9. Statistical Analysis

Statistical assays were carried out through the analysis of variance (ANOVA) test with SAS software (SAS Institute, Cary, NC, USA) and means were compared by the least significant difference (LSD) test or Tukey’s test dependent on the number of groups. Comparisons with p < 0.05 were considered significantly different.

3. Results

3.1. Waterlogging Tolerant Seedlings Exhibited Higher AOX Expression and Better Shoot Growth under Waterlogging

Here we identified an AOX gene (Cla97C10G192430) from the watermelon genome and named it as ClaAOX. Results from gene expression analysis showed that waterlogging upregulated the expression of ClaAOX in both cultivars, with the tolerant cultivar ‘YL’ showing significantly higher (p < 0.05) ClaAOX transcript level than the sensitive cultivar ‘Zaojia8424’ during 72 h of waterlogging stress (Figure 1A). For instance, ClaAOX expression at 24 h after stress (HAS) had increased by 201.1% and 510.7% in ‘Zaojia8424’ and ‘YL’, respectively, in comparison with the original level.
Shoot fresh weight was reduced by waterlogging, with a more significant (p < 0.05) decrease observed in ‘Zaojia8424’ (Figure 1B). A foliar spray of SHAM, an inhibitor of AOX, exacerbated the reduction of shoot fresh weight in both cultivars. However, there were no significant differences between ‘Zaojia8424’ and ‘YL’ in terms of shoot fresh weight under combined treatment of waterlogging and SHAM. Meanwhile, yellowing was observed in ‘Zaojia8424’ leaves but not in ‘YL’ leaves, when plants were exposed to waterlogging, and exogenous SHAM accelerated the wilting of both cultivars under waterlogging (Figure 1C).

3.2. Waterlogging and AOX Inhibition Reduced the Photosynthesis Performance in Both Cultivars

Photosynthesis is the main function of a leaf and is easily affected by environmental stresses such as waterlogging. Data in Figure 2 showed that the net photosynthesis rate (Pn), stomatal conductance (Gs) and transpiration rate (E) decreased with the prolonging of waterlogging in both cultivars. However, ‘YL’ was less affected than ‘Zaojia8424’ by waterlogging. For instance, Pn of ‘YL’ was approximately twofold that of ‘Zaojia8424’ after 6 days of waterlogging. Again, no significant differences were detected in Pn, Gs and E between cultivars in the combination treatment of waterlogging and SHAM (Figure 2A,B,D).
The intercellular CO2 concentration (Ci) was increased by waterlogging in both cultivars (Supplementary Figure S1). A more significant increase was observed in ‘Zaojia-8424’, compared with that in ‘YL’. However, no significant differences in Ci were detected between ‘Zaojia8424’ and ‘YL’ in waterlogging plus SHAM treatment (Figure 2C).
We found that Fv/Fm (maximum quantum efficiency of PSII photochemistry) and F’v/F’m (effective quantum use efficiency of PSII in the light-adapted state) was decreased significantly (p < 0.05) during waterlogging in ‘Zaojia8424’, whereas was not affected in ‘YL’ (Figure 3A,C). Exogenous SHAM decreased Fv/Fm and F’v/F’m in both cultivars under waterlogging (Figure 3A,C).
qP (proportion of open PSII) and YII (the effective quantum yield of PSII) was decreased by waterlogging in both cultivars, with a more profound reduction detected in ‘Zaojia8424’ (Figure 3B,D). YNPQ (the quantum yield of regulated non-photochemical energy loss in PSII) was enhanced by waterlogging in both cultivars, with more significant increase observed in ‘Zaojia8424’ (Figure 3E). YNO (the quantum yield of non-regulated energy loss in PSII) rose during waterlogging in ‘Zaojia8424’ rather than ‘YL’ (Figure 3F).
Interestingly, there were no significant differences in qP, F’v/F’m, YII, YNPQ, and YNO between ‘Zaojia8424’ and ‘YL’ in the combined treatment of waterlogging and SHAM.

3.3. Inhibition of AOX Activity Exacerbated the Oxidative Damage Induced by Waterlogging

Membrane permeability and lipid peroxidation are two typical markers of oxidative damage. Results in Figure 4 showed that electrolyte leakage and TBARS content increased significantly (p < 0.05) in both cultivars by waterlogging, with a more profound increase in the sensitive cultivar ‘Zaojia8424’ than the tolerant cultivar ‘YL’. In addition, the inhibition of AOX activity by SHAM aggravated the membrane damage, with no significant differences detected between two cultivars.

3.4. AOX Inhibition Promoted the ROS Accumulation in Watermelon Leaves during Waterlogging

The occurrence of oxidative damage is due to the excessive accumulation of ROS. Here, we investigated the effects of waterlogging and AOX inhibition on contents of O2− and H2O2 in watermelon seedlings. Results in Figure 5 showed that O2− and H2O2 contents increased during waterlogging in both cultivars, with less profound increases in O2− and H2O2 content observed in the tolerant cultivar ‘YL’. For instance, O2− content was increased by about 3 fold and 1 fold in ‘Zaojia8424’ and ‘YL’, respectively, after 9 days of waterlogging treatment. In addition, the inhibition of AOX activity by SHAM promoted further increases in O2− and H2O2 content in both cultivars, with no significant differences between cultivars (Figure 5A,B).
Superoxide are generated by the reduction of O2 by NADPH oxidase which is coded by the Rboh gene. Results from qPCR analysis showed that waterlogging upregulated the transcript levels of RbohA in both cultivars. However, no significant differences of RbohA expression were detected between ‘Zaojia8424’ and ‘YL’ (Figure 5C).

3.5. Responses of Antioxidant Enzymes to Waterlogging in Leaves

Antioxidant enzymes play crucial roles in the balance of ROS generation and scavenging. Here, we examined the activities of antioxidant enzymes under waterlogging in seedlings of both cultivars. Generally, the activity of SOD, CAT, APX, GPOD, DHAR and GR showed upward trends under waterlogging (Figure 6A–F). There were no significant differences in the activities of SOD, DHAR, and GR between ‘Zaojia8424’ and ‘YL’ (Figure 6A,E,F). However, the tolerant cultivar ‘YL’ exhibited a significantly (p < 0.05) higher CAT activity at 3 DAS and a significantly (p < 0.05) higher APX activity at 0 and 3 DAS (Figure 6B,D). Meanwhile, the increase of GPOD activity by waterlogging was more evident in ‘Zaojia8424’ than in ‘YL’ (Figure 6C).
Furthermore, we investigated the expressions of antioxidant genes. Generally, waterlogging induced the expression of CSD, CAT, APX, GPOD, DHAR, and GR in watermelon leaves (Figure 7A–F). There were no significant differences detected in CSD, DHAR and GR transcript levels between cultivars during 72 h of waterlogging (Figure 7A,E,F). The expressions of CAT and APX were significantly (p < 0.05) higher in the tolerant cultivar ‘YL’ than the sensitive cultivar ‘Zaojia8424’ (Figure 6B,D). The relative expression of GPOD was significantly (p < 0.05) higher in ‘Zaojia8424’ than in ‘YL’ during waterlogging (Figure 6C).

3.6. AOX Inhibition Hampered the Homeostasis of Glutathione and Ascorbate in Watermelon Seedlings under Waterlogging

Antioxidant substances are considered to be an important player during ROS scavenging. In the present study, we found that waterlogging decreased the contents of glutathione (GSH), oxidized glutathione (GSSG) and total glutathione in the seedlings of both cultivars (Figure 8A–C). Interestingly, the GSH to GSSG ratio increased significantly (p < 0.05) in the tolerant cultivar ‘YL’ during the second half of the 9 days of waterlogging, which showed a significantly higher level than the sensitive cultivar, ‘Zaojia8424’ (Figure 8D). The inhibition of AOX dramatically reduced the GSH/GSSG ratio with no significant differences detected between cultivars (Figure 8E).
The ascorbate acid (ASC) content in watermelon leaves during waterlogging was also investigated. Results showed that the ASC content increased initially, peaked at 6 DAS, and then declined (Figure 9A). The ASC content was significant higher (p < 0.05) in ‘YL’ than ‘Zaojia8424’ during 9 days of waterlogging treatment. For instance, ASC content was increased by 84.5% and 184.1% in ‘Zaojia8424’ and ‘YL’, respectively, compared with the original level. Additionally, the differences in ASC content between ‘Zaojia8424’ and ‘YL’ were diminished by AOX inhibition (Figure 9B).

4. Discussion

Alternative oxidase (AOX) has two subfamilies, AOX1 and AOX2. In general, AOX1 is induced by stresses, whereas AOX2 is thought to be developmentally expressed [40]. In the present study, we found that expression of ClaAOX was upregulated sharply in watermelon leaves of both cultivars. In our previous study, we have shown that activity of AOX was induced by waterlogging in watermelon roots [27]. These results indicated that ClaAOX might be waterlogging induced. Phylogenetic analysis suggested that ClaAOX was grouped in the AOX2 subfamily (Figure S2). Recently, some isoforms of AOX2 were experimentally validated to be responsive to abiotic stresses. For instance, CaAOX2A and CaAOX2D were upregulated by drought in legumes [41]. Moreover, tolerant cultivar “YL” displayed the higher level of ClaAOX than sensitive cultivar ’Zaojia8424′ in leaves during waterlogging stress (Figure 1A).
Photosynthesis inhibition is a typical phenomenon caused by waterlogging. In the present study, the waterlogging tolerant cultivar ‘YL’ exhibited higher Pn than the sensitive cultivar ‘Zaojia8424’ during waterlogging treatment (Figure 1A), which is consistent with previous results in sorghum [10], avocado (Persea americana) [42] and Actinidia valvata [43]. The tolerant cultivar of watermelon ‘YL’ displayed higher expression of ClaAOX compared with the sensitive cultivar ‘Zaojia8424’ under waterlogging (Figure 1A), and the inhibition of AOX activity diminished with the better performance of Pn in ‘YL’ under waterlogging (Figure 2A). Thus, it is suggested that the maintenance of photosynthesis under waterlogging in ‘YL’ was likely due, at least in part, to the AOX pathway. To the best of our knowledge, this is the first report about the involvement of AOX in optimizing photosynthesis under waterlogging. Similarly, it has been reported that inhibition of AOX activity by an exogenous inhibitor or AOX gene silencing reduced Pn when plants were exposed to drought [25], low light [44], low temperature and osmotic stress [45], and heat and high light [46] as well. However, overexpression of AOX increased Pn under drought [26]. Moreover, a single SHAM treatment did not affect the Pn in Pisum sativum under control conditions [45]. Thus, AOX might play a vital role in the maintenance of plant photosynthesis under abiotic stresses.
Reduced Pn by waterlogging is considered to be associated with the damage of the photosynthetic apparatus [47]. Fv/Fm, an indicator of the photoinhibition of PSII [48], decreased to about 0.75 by waterlogging in ‘Zaojia8424’ (Figure 4A), suggesting the occurrence of photoinhibition. Similar phenomena were observed in Arabidopsis [49], sorghum [10], tomato [11] and Jerusalem artichoke (Helianthus tuberosus) [12], However, while Fv/Fm was hardly affected in ‘YL’ by waterlogging, it was decreased sharply by AOX inhibition, suggesting that AOX is likely involved in the maintenance of PSII function under waterlogging. Similarly, Fv/Fm was decreased by severe drought in AOX knockdown plants [26].
The energy absorbed by PSII can be divided into three partitions, the quantum yield of photosynthesis (YII), the quantum yield of regulated non-photochemical energy loss (YNPQ) and the quantum yield of non-regulated energy loss (YNO), according Kramer et al. [50]. Here we found that YII was reduced by waterlogging in both cultivars (Figure 4B), which is consistent with previous studies [10,49]. Moreover, the decrease of YII resulted from the decline of qP and Fv’/Fm’, since YII is a product of qP and Fv’/Fm’ [51]. Besides, ‘YL’ exhibited significantly (p < 0.05) higher YII than ‘Zaojia8424’, but this effect was reduced by AOX inhibition (Figure 4B). Decreased YII by SHAM implies lower electron transport to carbon fixation [21], which was correlated to the reduced Pn (Figure 2A).
The increase of YNO indicates the insufficiency of photochemical energy conversion and photo-protective regulatory mechanism, and the excess electron can be used to form singlet oxygen (1O2) in the chloroplast [52]. YNO was increased by SHAM in ‘YL’, suggesting inhibition of AOX could result in more ROS generation in the chloroplast. Since AOX may act as an additional electron sink for photo-generated reductant [25], the inhibition of AOX activity would weaken the sink, and thus impair the photosynthetic capacity.
Oxidative damage occurred when plants are exposed to long term waterlogging [53]. Here, watermelon seedlings suffered from oxidative stress, since increased O2− and H2O2 content and consequently higher TBARS content and electrolyte leakage were detected in both cultivars after waterlogging treatment (Figure 4A,B and Figure 5A,B). The tolerant cultivar ‘YL’, with higher transcript levels of ClaAOX, displayed less severe oxidative damage, compared with the sensitive cultivar ‘Zaojia8424’. However, this alleviation of oxidative damage in ‘YL’ was reduced by SHAM. Similar phenomena were observed in the roots of these two cultivars in our previous study [27]. Other studies have reported that overexpression of AOX decreased TBARS contents of tobacco under hypoxia, and knockdown of AOX increased TBARS contents [24]. In addition, AOX knockdown plants or mutants had increased contents of H2O2 when plants were exposed to drought stress [54], arsenic stress [55], nitrogen deficiency and iron deficiency [46]. Overexpression of AOX decreased the H2O2 content under salt stress [56] and drought stress [54]. Thus, AOX might be essential in maintaining plant ROS balance under abiotic stresses.
AOX may ameliorate oxidative stress via two pathways, modulating the production of ROS and inducing the antioxidant capacity. For instance, Maxwell et al. [22] reported that AOX antisense tobacco cells exhibited higher levels of ROS, whereas AOX overexpression cells displayed lower ROS abundance compared with wild-type cells. They further found that AOX overexpression cells had lower expression of genes encoding antioxidant enzymes, such as SODA, SODB, CAT and GPX [22]. Thus, AOX has been considered as a key a player in reducing mitochondrial ROS formation in plant cells.
However, the cells they used lacked developed chloroplasts [22]. The responses of the antioxidant system might be different from tissues containing developed chloroplasts. For instance, we found that expression of CSD, CAT, APX, DHAR and GR were upregulated by waterlogging in watermelon leaves (Figure 7), whereas the expressions of the corresponding genes were downregulated sharply in watermelon roots in our previous study using the same cultivars [27]. These different antioxidant responses to hypoxia between shoot and roots were observed in barley [18] and gray poplar (Populus × canescens) [3]. Skutnik and Rychter [18] further suggested that AOX plays a major role in ROS scavenging in roots, whereas antioxidant substances including glutathione and ascorbate are of great importance in the ROS detoxification in leaves.
Thus, the responses of the glutathione ascorbate cycle in watermelon leaves under waterlogging were further examined, where we found that GSH, GSSG and total glutathione contents were decreased by waterlogging (Figure 8A–C). As GSH regenerates ascorbate by reducing DHA through the glutathione-ascorbate cycle [57], the reduced GSH could be attributed to the increase of ASC content under waterlogging (Figure 9A). In addition, the tolerant cultivar ‘YL’, with a higher expression level of ClaAOX, exhibited higher ASC contents compared with the sensitive cultivar ‘Zaojia8424’. However, this effect was abolished by AOX inhibition, implying the regulation of the glutathione-ascorbate cycle by AOX. To the best of our knowledge, this is the first report of the involvement of the glutathione-ascorbate cycle in antioxidant responses mediated by AOX under waterlogging. Since ASC is the substrate of APX, a higher content of ASC in ‘YL’ resulted in enhanced activity of APX and transcript level of APX (Figure 6D and Figure 7D), which is beneficial for ROS scavenging.
Interestingly, the GSH/GSSG ratio was increased in the tolerant cultivar ‘YL’, but not in the sensitive cultivar ‘Zaojia8424’ (Figure 8D). However, the increased GSH/GSSG ratio in ‘YL’ was hampered by AOX inhibition (Figure 8E). Similarly, it has been reported that inhibition of AOX by SHAM or AOX gene knockdown decreased the GSH/GSSG ratio when plants were exposed to As stress [55] and low temperature [48]. The GSH/GSSG ratio, presenting the redox state of cells, has long been recognized as a signaling cascade [58]. Since CAT is sensitive to glutathione redox [59], the higher GSH/GSSG ratio in ‘YL’ resulted in the higher activity of CAT and increased expression of CAT compared with ‘Zaojia8424’ (Figure 6B and Figure 7B). Conversely, GPOD is considered to be a biomarker of oxidative stress [60], and thus the lower GSH/GSSG ratio in ‘Zaojia8424’ would lead to higher activity of GPOD and increased expression of GPOD compared with ‘YL’ (Figure 6C and Figure 7C).

5. Conclusions

Waterlogging decreased leaf Pn and YII in watermelon. The photosynthesis was less affected in the waterlogging tolerant cultivar ‘YL’ than the sensitive cultivar ‘Zaojia8424’. Moreover, waterlogging induced photoinhibition in ‘Zaojia8424’ but not in ‘YL’. Meanwhile, waterlogging promoted the accumulation of ROS, and triggered oxidative damage. The waterlogging tolerant cultivar ‘YL’ suffered from less severe oxidative damage due to increased contents of ascorbate, a higher ratio of GSH/GSSG, a higher activity of APX and CAT, and enhanced transcript levels of CAT and APX compared to the sensitive cultivar ‘Zaojia8424’. However, no significant differences were detected in photosynthesis and antioxidant performances between cultivars when AOX activity was inhibited by SHAM application. Our results address the involvement of photosynthetic electronic transfer and glutathione-ascorbate cycle in AOX manipulated waterlogging tolerance in watermelon seedlings.

Supplementary Materials

The following are available online at https://www.mdpi.com/article/10.3390/horticulturae7060130/s1, Figure S1: Phylogenetic analysis of the alternative oxidase (AOX) amino acid sequences from watermelon genome; Table S1: Primer sequences used in the qRT-PCR experiments.

Author Contributions

Conceptualization, Z.Z. and Y.H. (Yong He); validation, C.F., J.Z., N.S. and Y.H. (Yunping Huang); investigation, J.Z., C.F. and N.S.; resources, Y.W. and Y.H. (Yunping Huang); writing—original draft preparation, Y.H. (Yong He) and Z.Z.; writing—review and editing, L.R. and Y.L.; project administration, Y.H. (Yong He) and Z.Z.; funding acquisition, Y.H. (Yong He) and Y.W. All authors have read and agreed to the published version of the manuscript.

Funding

This research was funded by National Natural Science Foundation of China (32072556), the key R&D Program of Zhejiang Province (No. 2019C02012), and the Major Science and Technology Project of Ningbo (2019B10007).

Institutional Review Board Statement

Not applicable.

Informed Consent Statement

Not applicable.

Data Availability Statement

Not applicable.

Conflicts of Interest

The authors declare no conflict of interest.

References

  1. Ahsan, N.; Lee, D.G.; Lee, S.H.; Lee, K.W.; Bahk, J.D.; Lee, B.H. A proteomic screen and identification of waterlogging-regulated proteins in tomato roots. Plant Soil 2007, 295, 37–51. [Google Scholar] [CrossRef]
  2. Fukao, T.; Barrera-Figueroa, B.E.; Juntawong, P.; Peña-Castro, J.M. Submergence and waterlogging stress in plants: A review highlighting research opportunities and understudied aspects. Front. Plant Sci. 2019, 10, 340. [Google Scholar] [CrossRef]
  3. Kreuzwieser, J.; Hauberg, J.; Howell, K.A.; Carroll, A.; Rennenberg, H.; Millar, A.H.; Whelan, J. Differential response of gray poplar leaves and roots underpins stress adaptation during hypoxia. Plant Physiol. 2009, 149, 461–473. [Google Scholar] [CrossRef] [PubMed] [Green Version]
  4. Lee, Y.H.; Kim, K.S.; Jang, Y.S.; Hwang, J.H.; Lee, D.H.; Choi, I.H. Global gene expression responses to waterlogging in leaves of rape seedlings. Plant Cell Rep. 2014, 33, 289–299. [Google Scholar] [CrossRef] [PubMed]
  5. Loreti, E.; van Veen, H.; Perata, P. Plant responses to flooding stress. Curr. Opin. Plant Biol. 2016, 33, 64–71. [Google Scholar] [CrossRef] [Green Version]
  6. De San Celedonio, R.P.; Abeledo, L.G.; Mantese, A.I.; Miralles, D.J. Differential root and shoot biomass recovery in wheat and barley with transient waterlogging during preflowering. Plant Soil 2017, 417, 481–498. [Google Scholar] [CrossRef]
  7. Barickman, T.C.; Simpson, C.R.; Sams, C.E. Waterlogging causes early modification in the physiological performance, carotenoids, chlorophylls, proline, and soluble sugars of cucumber plants. Plants 2019, 8, 160. [Google Scholar] [CrossRef] [Green Version]
  8. Salah, A.; Zhan, M.; Cao, C.; Han, Y.; Ling, L.; Liu, Z.; Li, P.; Ye, M.; Jiang, Y. Gamma-aminobutyric acid promotes chloroplast ultrastructure, antioxidant capacity, and growth of waterlogged maize seedlings. Sci. Rep. 2019, 9, 484. [Google Scholar] [CrossRef] [PubMed]
  9. Zeng, R.; Chen, L.; Wang, X.; Cao, J.; Li, X.; Xu, X.; Xia, Q.; Chen, T.; Zhang, L. Effect of waterlogging stress on dry matter accumulation, photosynthesis characteristics, yield, and yield components in three different ecotypes of peanut (Arachis hypogaea L.). Agronomy 2020, 10, 1244. [Google Scholar] [CrossRef]
  10. Zhang, R.D.; Zhou, Y.F.; Yue, Z.X.; Chen, X.F.; Cao, X.; Xu, X.X.; Xing, Y.F.; Jiang, B.; Ai, X.Y.; Huang, R.D. Changes in photosynthesis, chloroplast ultrastructure, and antioxidant metabolism in leaves of sorghum under waterlogging stress. Photosynthetica 2019, 57, 1076–1083. [Google Scholar] [CrossRef] [Green Version]
  11. Mauro, R.P.; Agnello, M.; Distefano, M.; Sabatino, L.; San Bautista Primo, A.; Leonardi, C.; Giuffrida, F. Chlorophyll fluorescence, photosynthesis and growth of tomato plants as affected by long-term oxygen root zone deprivation and grafting. Agronomy 2020, 10, 137. [Google Scholar] [CrossRef] [Green Version]
  12. Yan, K.; Zhao, S.; Cui, M.; Han, G.; Wen, P. Vulnerability of photosynthesis and photosystem I in Jerusalem artichoke (Helianthus tuberosus L.) exposed to waterlogging. Plant Physiol. Biochem. 2018, 125, 239–246. [Google Scholar] [CrossRef]
  13. Ishida, S.; Uebayashi, N.; Tazoe, Y.; Ikeuchi, M.; Homma, K.; Sato, F.; Endo, T. Diurnal and developmental changes in energy allocation of absorbed light at PSII in field-grown rice. Plant Cell Physiol. 2014, 55, 171–182. [Google Scholar] [CrossRef] [Green Version]
  14. Mittler, R. Oxidative stress, antioxidants and stress tolerance. Trends Plant Sci. 2002, 7. [Google Scholar] [CrossRef]
  15. Noctor, G.; Reichheld, J.P.; Foyer, C.H. ROS-related redox regulation and signaling in plants. Semin. Cell Dev. Biol. 2018, 80, 3–12. [Google Scholar] [CrossRef] [Green Version]
  16. Sachdev, S.; Ansari, S.A.; Ansari, M.I.; Fujita, M.; Hasanuzzaman, M. Abiotic stress and reactive oxygen species: Generation, signaling, and defense mechanisms. Antioxidants 2021, 10, 277. [Google Scholar] [CrossRef] [PubMed]
  17. Zheng, C.; Jiang, D.; Liu, F.; Dai, T.; Jing, Q.; Cao, W. Effects of salt and waterlogging stresses and their combination on leaf photosynthesis, chloroplast ATP synthesis, and antioxidant capacity in wheat. Plant Sci. 2009, 176, 575–582. [Google Scholar] [CrossRef] [PubMed]
  18. Skutnik, M.; Rychter, A.M. Differential response of antioxidant systems in leaves and roots of barley subjected to anoxia and post-anoxia. J. Plant Physiol. 2009, 166, 926–937. [Google Scholar] [CrossRef] [PubMed]
  19. Apel, K.; Hirt, H. Reactive oxygen species: Metabolism, oxidative stress, and signal transduction. Annu. Rev. Plant Biol. 2004, 55, 373–399. [Google Scholar] [CrossRef] [Green Version]
  20. Noctor, G.; Foyer, C.H. Ascorbate and glutathione: Keeping active oxygen under control. Annu. Rev. Plant Biol. 1998, 49, 249–279. [Google Scholar] [CrossRef]
  21. Hasanuzzaman, M.; Bhuyan, M.; Anee, T.I.; Parvin, K.; Nahar, K.; Al Mahmud, J.; Fujita, M. Regulation of ascorbate-glutathione pathway in mitigating oxidative damage in plants under abiotic stress. Antioxidants 2019, 8, 384. [Google Scholar] [CrossRef] [Green Version]
  22. Maxwell, D.P.; Wang, Y.; McIntosh, L. The alternative oxidase lowers mitochondrial reactive oxygen production in plant cells. Proc. Natl. Acad. Sci. USA 1999, 96, 8271–8276. [Google Scholar] [CrossRef] [Green Version]
  23. Selinski, J.; Scheibe, R.; Day, D.A.; Whelan, J. alternative oxidase is positive for plant performance. Trends Plant Sci. 2018, 23, 588–597. [Google Scholar] [CrossRef]
  24. Jayawardhane, J.; Cochrane, D.W.; Vyas, P.; Bykova, N.V.; Vanlerberghe, G.C.; Igamberdiev, A.U. Roles for plant mitochondrial alternative oxidase under normoxia, hypoxia, and reoxygenation conditions. Front. Plant Sci. 2020, 11, 566. [Google Scholar] [CrossRef]
  25. Dahal, K.; Wang, J.; Martyn, G.D.; Rahimy, F.; Vanlerberghe, G.C. Mitochondrial alternative oxidase maintains respiration and preserves photosynthetic capacity during moderate drought in Nicotiana tabacum. Plant Physiol. 2014, 166, 1560–1574. [Google Scholar] [CrossRef] [PubMed] [Green Version]
  26. Dahal, K.; Vanlerberghe, G.C. Alternative oxidase respiration maintains both mitochondrial and chloroplast function during drought. New Phytol. 2017, 213, 560–571. [Google Scholar] [CrossRef] [PubMed]
  27. Zheng, J.; Ying, Q.; Fang, C.; Sun, N.; Si, M.; Yang, J.; Zhu, B.; Ruan, Y.-L.; Zhu, Z.; He, Y. Alternative oxidase pathway is likely involved in waterlogging tolerance of watermelon. Sci. Hortic. 2021, 278, 109831. [Google Scholar] [CrossRef]
  28. Dionisio-Sese, M.L.; Tobita, S. Antioxidant responses of rice seedlings to salinity stress. Plant Sci. 1998, 135, 1–9. [Google Scholar] [CrossRef]
  29. Cakmak, I.; Marschner, H. Magnesium deficiency and high light intensity enhance activities of superoxide dismutase, ascorbate peroxidase, and glutathione reductase in bean leaves. Plant Physiol. 1992, 98, 1222–1227. [Google Scholar] [CrossRef] [Green Version]
  30. He, Y.; Ye, Z.X.; Ying, Q.S.; Ma, Y.P.; Zang, Y.X.; Wang, H.S.; Yu, Y.J.; Zhu, Z.J. Glyoxylate cycle and reactive oxygen species metabolism are involved in the improvement of seed vigor in watermelon by exogenous GA(3). Sci. Hortic. 2019, 247, 184–194. [Google Scholar] [CrossRef]
  31. Cheeseman, J.M. Hydrogen peroxide concentrations in leaves under natural conditions. J. Exp. Bot. 2006, 57, 2435–2444. [Google Scholar] [CrossRef] [PubMed] [Green Version]
  32. He, Y.; Zhu, Z.; Yang, J.; Ni, X.; Zhu, B. Grafting increases the salt tolerance of tomato by improvement of photosynthesis and enhancement of antioxidant enzymes activity. Environ. Exp. Bot. 2009, 66, 270–278. [Google Scholar] [CrossRef]
  33. Rao, K.M.; Sresty, T. Antioxidative parameters in the seedlings of pigeonpea (Cajanus cajan (L.) Millspaugh) in response to Zn and Ni stresses. Plant Sci. 2000, 157, 113–128. [Google Scholar] [CrossRef]
  34. Egley, G.; Paul, R.; Vaughn, K.; Duke, S. Role of peroxidase in the development of water-impermeable seed coats in Sida spinosa L. Planta 1983, 157, 224–232. [Google Scholar] [CrossRef]
  35. Nakano, Y.; Asada, K. Hydrogen peroxide is scavenged by ascorbate-specific peroxidase in spinach chloroplasts. Plant Cell Physiol. 1981, 22, 867–880. [Google Scholar] [CrossRef]
  36. Foyer, C.H.; Halliwell, B. The presence of glutathione and glutathione reductase in chloroplasts: A proposed role in ascorbic acid metabolism. Planta 1976, 133, 21–25. [Google Scholar] [CrossRef]
  37. Hodges, D.M.; Andrews, C.J.; Johnson, D.A.; Hamilton, R.I. Antioxidant compound responses to chilling stress in differentially sensitive inbred maize lines. Physiol. Plant. 1996, 98, 685–692. [Google Scholar] [CrossRef]
  38. Li, H.; Jiang, X.; Lv, X.; Ahammed, G.J.; Guo, Z.; Qi, Z.; Yu, J.; Zhou, Y. Tomato GLR3.3 and GLR3.5 mediate cold acclimation-induced chilling tolerance by regulating apoplastic H2O2 production and redox homeostasis. Plant Cell Environ. 2019, 42, 3326–3339. [Google Scholar] [CrossRef]
  39. Livak, K.J.; Schmittgen, T.D. Analysis of relative gene expression data using real-time quantitative PCR and the 2−ΔΔCT method. Methods 2001, 25, 402–408. [Google Scholar] [CrossRef]
  40. Considine, M.J.; Holtzapffel, R.C.; Day, D.A.; Whelan, J.; Millar, A.H. Molecular distinction between alternative oxidase from monocots and dicots. Plant Physiol. 2002, 129, 949–953. [Google Scholar] [CrossRef] [Green Version]
  41. Sweetman, C.; Soole, K.L.; Jenkins, C.L.; Day, D.A. Genomic structure and expression of alternative oxidase genes in legumes. Plant Cell Environ. 2019, 42, 71–84. [Google Scholar] [CrossRef] [Green Version]
  42. Doupis, G.; Kavroulakis, N.; Psarras, G.; Papadakis, I.E. Growth, photosynthetic performance and antioxidative response of ‘Hass’ and ‘Fuerte’ avocado (Persea americana Mill.) plants grown under high soil moisture. Photosynthetica 2017, 55, 655–663. [Google Scholar] [CrossRef]
  43. Li, Z.; Zhong, Y.; Bai, D.; Lin, M.; Qi, X.; Fang, J. Comparative analysis of physiological traits of three Actinidia valvata Dunn genotypes during waterlogging and post-waterlogging recovery. Hortic. Environ. Biotechnol. 2020, 61, 825–836. [Google Scholar] [CrossRef]
  44. Yoshida, K.; Terashima, I.; Noguchi, K. Distinct roles of the cytochrome pathway and alternative oxidase in leaf photosynthesis. Plant Cell Physiol. 2006, 47, 22–31. [Google Scholar] [CrossRef]
  45. Dinakar, C.; Vishwakarma, A.; Raghavendra, A.S.; Padmasree, K. Alternative oxidase pathway optimizes photosynthesis during osmotic and temperature stress by regulating cellular ROS, malate valve and antioxidative systems. Front. Plant Sci. 2016, 7, 68. [Google Scholar] [CrossRef] [PubMed] [Green Version]
  46. Murik, O.; Tirichine, L.; Prihoda, J.; Thomas, Y.; Araujo, W.L.; Allen, A.E.; Fernie, A.R.; Bowler, C. Downregulation of mitochondrial alternative oxidase affects chloroplast function, redox status and stress response in a marine diatom. New Phytol. 2019, 221, 1303–1316. [Google Scholar] [CrossRef] [Green Version]
  47. Striker, G.G.; Colmer, T.D. Flooding tolerance of forage legumes. J. Exp. Bot. 2017, 68, 1851–1872. [Google Scholar] [CrossRef] [PubMed] [Green Version]
  48. Aro, E.-M.; Virgin, I.; Andersson, B. Photoinhibition of photosystem II. Inactivation, protein damage and turnover. Biochim. Biophys. Acta Bioenerg. 1993, 1143, 113–134. [Google Scholar] [CrossRef]
  49. Xu, L.; Pan, R.; Shabala, L.; Shabala, S.; Zhang, W.-Y. Temperature influences waterlogging stress-induced damage in Arabidopsis through the regulation of photosynthesis and hypoxia-related genes. Plant Growth Regul. 2019, 89, 143–152. [Google Scholar] [CrossRef]
  50. Kramer, D.M.; Johnson, G.; Kiirats, O.; Edwards, G.E. New fluorescence parameters for the determination of QA redox state and excitation energy fluxes. Photosynth. Res. 2004, 79, 209–218. [Google Scholar] [CrossRef]
  51. Genty, B.; Briantais, J.-M.; Baker, N.R. The relationship between the quantum yield of photosynthetic electron transport and quenching of chlorophyll fluorescence. Biochim. Biophys. Acta Gen. Subj. 1989, 990, 87–92. [Google Scholar] [CrossRef]
  52. Baycu, G.; Moustaka, J.; Gevrek, N.; Moustakas, M. Chlorophyll fluorescence imaging analysis for elucidating the mechanism of photosystem II acclimation to cadmium exposure in the hyperaccumulating plant Noccaea caerulescens. Materials 2018, 11, 2580. [Google Scholar] [CrossRef] [Green Version]
  53. Xiao, Y.; Wu, X.; Sun, M.; Peng, F. Hydrogen sulfide alleviates waterlogging-induced damage in peach seedlings via enhancing antioxidative system and inhibiting ethylene synthesis. Front. Plant Sci. 2020, 11, 696. [Google Scholar] [CrossRef] [PubMed]
  54. Zhu, T.; Zou, L.; Li, Y.; Yao, X.; Xu, F.; Deng, X.; Zhang, D.; Lin, H. Mitochondrial alternative oxidase-dependent autophagy involved in ethylene-mediated drought tolerance in Solanum lycopersicum. Plant Biotech. J. 2018, 16, 2063–2076. [Google Scholar] [CrossRef] [Green Version]
  55. Demircan, N.; Cucun, G.; Uzilday, B. Mitochondrial alternative oxidase (AOX1a) is required for the mitigation of arsenic-induced oxidative stress in Arabidopsis thaliana. Plant Biotechnol. Rep. 2020, 14, 235–245. [Google Scholar] [CrossRef]
  56. Smith, C.A.; Melino, V.J.; Sweetman, C.; Soole, K.L. Manipulation of alternative oxidase can influence salt tolerance in Arabidopsis thaliana. Physiol. Plant. 2009, 137, 459–472. [Google Scholar] [CrossRef]
  57. Foyer, C.H.; Noctor, G. Ascorbate and glutathione: The heart of the redox hub. Plant Physiol. 2011, 155, 2–18. [Google Scholar] [CrossRef] [Green Version]
  58. May, M.J.; Vernoux, T.; Leaver, C.; Montagu, M.V.; Inze, D. Glutathione homeostasis in plants: Implications for environmental sensing and plant development. J. Exp. Bot. 1998, 49, 649–667. [Google Scholar] [CrossRef] [Green Version]
  59. Sharma, P.; Jha, A.B.; Dubey, R.S.; Pessarakli, M. Reactive oxygen species, oxidative damage, and antioxidative defense mechanism in plants under stressful conditions. J. Bot. 2012, 2012, 1–26. [Google Scholar] [CrossRef] [Green Version]
  60. Jouili, H.; Bouazizi, H.; El Ferjani, E. Plant peroxidases: Biomarkers of metallic stress. Acta Physiol. Plant. 2011, 33, 2075–2082. [Google Scholar] [CrossRef]
Horticulturae 07 00130 g001 550
Figure 1. Changes of ClaAOX expression in watermelon leaves under waterlogging (A) and effects of SHAM on plant growth in two cultivars of watermelon (‘Zaojia8424’ and ‘YL’) under waterlogging ((B) shoot fresh weight. (C) Typical leaf phenotype under waterlogging and SHAM). Each histogram represents a mean ± SE of four replicates (n = 4). Different letters indicate significant differences between treatments (p < 0.05) according to Tukey’s test and * indicates a significant difference for ‘YL’ compared to ‘Zaojia8424’ at p < 0.05 according to the least significant difference (LSD) test.
Figure 1. Changes of ClaAOX expression in watermelon leaves under waterlogging (A) and effects of SHAM on plant growth in two cultivars of watermelon (‘Zaojia8424’ and ‘YL’) under waterlogging ((B) shoot fresh weight. (C) Typical leaf phenotype under waterlogging and SHAM). Each histogram represents a mean ± SE of four replicates (n = 4). Different letters indicate significant differences between treatments (p < 0.05) according to Tukey’s test and * indicates a significant difference for ‘YL’ compared to ‘Zaojia8424’ at p < 0.05 according to the least significant difference (LSD) test.
Horticulturae 07 00130 g001
Horticulturae 07 00130 g002 550
Figure 2. Effects of exogenous SHAM on photosynthesis in watermelon leaves under waterlogging treatment. (A) Net photosynthesis rate (Pn). (B) Stomatal conductance (Gs). (C) Intercellular CO2 concentration (Ci). (D) Transpiration rate (E). Different letters indicate significant differences according to Tukey’s test (p < 0.05).
Figure 2. Effects of exogenous SHAM on photosynthesis in watermelon leaves under waterlogging treatment. (A) Net photosynthesis rate (Pn). (B) Stomatal conductance (Gs). (C) Intercellular CO2 concentration (Ci). (D) Transpiration rate (E). Different letters indicate significant differences according to Tukey’s test (p < 0.05).
Horticulturae 07 00130 g002
Horticulturae 07 00130 g003 550
Figure 3. Effects of exogenous SHAM on chlorophyll fluorescence parameters in watermelon leaves under waterlogging treatment. (A) Fv/Fm (maximum quantum efficiency of PSII photochemistry). (B) qP (proportion of open PSII). (C) F’v/F’m (effective quantum use efficiency of PSII in the light-adapted state). (D) YII (the effective quantum yield of PSII). (E) YNPQ (the quantum yield of regulated non-photochemical energy loss in PSII). (F) YNO (the quantum yield of non-regulated energy loss in PSII). Different letters indicate significant differences according to Tukey’s test (p < 0.05).
Figure 3. Effects of exogenous SHAM on chlorophyll fluorescence parameters in watermelon leaves under waterlogging treatment. (A) Fv/Fm (maximum quantum efficiency of PSII photochemistry). (B) qP (proportion of open PSII). (C) F’v/F’m (effective quantum use efficiency of PSII in the light-adapted state). (D) YII (the effective quantum yield of PSII). (E) YNPQ (the quantum yield of regulated non-photochemical energy loss in PSII). (F) YNO (the quantum yield of non-regulated energy loss in PSII). Different letters indicate significant differences according to Tukey’s test (p < 0.05).
Horticulturae 07 00130 g003
Horticulturae 07 00130 g004 550
Figure 4. Effects of exogenous SHAM on oxidative damage in watermelon leaves under waterlogging treatment. (A) Electrolyte leakage. (B) TBARS content. Different letters indicate significant differences according to Tukey’s test (p < 0.05).
Figure 4. Effects of exogenous SHAM on oxidative damage in watermelon leaves under waterlogging treatment. (A) Electrolyte leakage. (B) TBARS content. Different letters indicate significant differences according to Tukey’s test (p < 0.05).
Horticulturae 07 00130 g004
Horticulturae 07 00130 g005 550
Figure 5. Effects of exogenous SHAM on ROS content in watermelon leaves under waterlogging treatment ((A), O2− content. (B), H2O2 content) and changes of RbohA expression in watermelon leaves during waterlogging (C). Different letters indicate significant differences according to Tukey’s test and * indicates a significant difference for ‘YL’ compared to ‘Zaojia8424’ at p < 0.05 (n = 4) according to LSD test.
Figure 5. Effects of exogenous SHAM on ROS content in watermelon leaves under waterlogging treatment ((A), O2− content. (B), H2O2 content) and changes of RbohA expression in watermelon leaves during waterlogging (C). Different letters indicate significant differences according to Tukey’s test and * indicates a significant difference for ‘YL’ compared to ‘Zaojia8424’ at p < 0.05 (n = 4) according to LSD test.
Horticulturae 07 00130 g005
Horticulturae 07 00130 g006 550
Figure 6. Effects of waterlogging on activities of antioxidant enzymes in watermelon. (A) SOD. (B) CAT. (C) GPOD. (D) APX. (E) DHAR. (F) GR. * indicates a significant difference for ‘YL’ compared to ‘Zaojia8424’ at p < 0.05 according to LSD test.
Figure 6. Effects of waterlogging on activities of antioxidant enzymes in watermelon. (A) SOD. (B) CAT. (C) GPOD. (D) APX. (E) DHAR. (F) GR. * indicates a significant difference for ‘YL’ compared to ‘Zaojia8424’ at p < 0.05 according to LSD test.
Horticulturae 07 00130 g006
Horticulturae 07 00130 g007 550
Figure 7. Effects of waterlogging on expressions of antioxidant genes in watermelon. (A) CSD. (B) CAT. (C) GPOD. (D) APX. (E) DHAR. (F) GR. * indicates a significant difference for ‘YL’ compared to ‘Zaojia8424’ at p < 0.05 according to LSD test.
Figure 7. Effects of waterlogging on expressions of antioxidant genes in watermelon. (A) CSD. (B) CAT. (C) GPOD. (D) APX. (E) DHAR. (F) GR. * indicates a significant difference for ‘YL’ compared to ‘Zaojia8424’ at p < 0.05 according to LSD test.
Horticulturae 07 00130 g007
Horticulturae 07 00130 g008 550
Figure 8. Changes of glutathione contents in watermelon leaves during waterlogging and effects of SHAM on GSH/GSSG ratio under waterlogging. (A) Total glutathione content under waterlogging. (B) GSH content under waterlogging. (C) GSSG content under waterlogging. (D) GSH/GSSG ratio during waterlogging. (E) Effects of SHAM on GSH/GSSG ratio under waterlogging. Different letters indicate significant differences according to Tukey’s test and * indicates a significant difference for ‘YL’ compared to ‘Zaojia8424’ at p < 0.05 (n = 4) according to LSD test.
Figure 8. Changes of glutathione contents in watermelon leaves during waterlogging and effects of SHAM on GSH/GSSG ratio under waterlogging. (A) Total glutathione content under waterlogging. (B) GSH content under waterlogging. (C) GSSG content under waterlogging. (D) GSH/GSSG ratio during waterlogging. (E) Effects of SHAM on GSH/GSSG ratio under waterlogging. Different letters indicate significant differences according to Tukey’s test and * indicates a significant difference for ‘YL’ compared to ‘Zaojia8424’ at p < 0.05 (n = 4) according to LSD test.
Horticulturae 07 00130 g008
Horticulturae 07 00130 g009 550
Figure 9. Changes of ASC contents in watermelon leaves during waterlogging (A) and effects of SHAM on ASC contents in watermelon leaves under waterlogging (B). Different letters indicate significant differences according to Tukey’s test and * indicates a significant difference for ‘YL’ compared to ‘Zaojia8424’ at p < 0.05 (n = 4) according to LSD test.
Figure 9. Changes of ASC contents in watermelon leaves during waterlogging (A) and effects of SHAM on ASC contents in watermelon leaves under waterlogging (B). Different letters indicate significant differences according to Tukey’s test and * indicates a significant difference for ‘YL’ compared to ‘Zaojia8424’ at p < 0.05 (n = 4) according to LSD test.
Horticulturae 07 00130 g009
Publisher’s Note: MDPI stays neutral with regard to jurisdictional claims in published maps and institutional affiliations.

Share and Cite

MDPI and ACS Style

Zheng, J.; Fang, C.; Ru, L.; Sun, N.; Liu, Y.; Huang, Y.; Wang, Y.; Zhu, Z.; He, Y. Role of Glutathione-Ascorbate Cycle and Photosynthetic Electronic Transfer in Alternative Oxidase-Manipulated Waterlogging Tolerance in Watermelon Seedlings. Horticulturae 2021, 7, 130. https://doi.org/10.3390/horticulturae7060130

AMA Style

Zheng J, Fang C, Ru L, Sun N, Liu Y, Huang Y, Wang Y, Zhu Z, He Y. Role of Glutathione-Ascorbate Cycle and Photosynthetic Electronic Transfer in Alternative Oxidase-Manipulated Waterlogging Tolerance in Watermelon Seedlings. Horticulturae. 2021; 7(6):130. https://doi.org/10.3390/horticulturae7060130

Chicago/Turabian Style

Zheng, Jiawen, Chunying Fang, Lei Ru, Nan Sun, Yuanyuan Liu, Yunping Huang, Yuhong Wang, Zhujun Zhu, and Yong He. 2021. "Role of Glutathione-Ascorbate Cycle and Photosynthetic Electronic Transfer in Alternative Oxidase-Manipulated Waterlogging Tolerance in Watermelon Seedlings" Horticulturae 7, no. 6: 130. https://doi.org/10.3390/horticulturae7060130

Note that from the first issue of 2016, this journal uses article numbers instead of page numbers. See further details here.

Article Metrics

Back to TopTop