Cucumber Phospholipase D alpha gene overexpression in tobacco enhanced drought stress tolerance by regulating stomatal closure and lipid peroxidation
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Plant phospholipase D (PLD), which can hydrolyze membrane phospholipids to produce phosphatidic acid (PA), a secondary signaling molecule, has been proposed to function in diverse plant stress responses. Both PLD and PA play key roles in plant growth, development, and cellular processes. PLD was suggested to mediate the regulation of stomatal movements by abscisic acid (ABA) as a response to water deficit. In this research, we characterized the roles of the cucumber phospholipase D alpha gene (CsPLDα, GenBank accession number EF363796) in the growth and tolerance of transgenic tobacco (Nicotiana tabacum) to drought stress.
The CsPLDα overexpression in tobacco lines correlated with the ABA synthesis and metabolism, regulated the rapid stomatal closure in drought stress, and reduced the water loss. The NtNCED1 expression levels in the transgenic lines and wild type (WT) were sharply up-regulated after 16 days of drought stress compared with those before treatment, and the expression level in the transgenic lines was significantly higher than that in the WT. The NtAOG expression level evidently improved after 8 and 16 days compared with that at 0 day of treatment and was significantly lower in the transgenic lines than in the WT. The ABA content in the transgenic lines was significantly higher than that in the WT. The CsPLDα overexpression could increase the osmolyte content and reduce the ion leakage. The proline, soluble sugar, and soluble protein contents significantly increased. By contrast, the electrolytic leakage and malondialdehyde accumulation in leaves significantly decreased. The shoot and root fresh and dry weights of the overexpression lines significantly increased. These results indicated that a significant correlation between CsPLDα overexpression and improved resistance to water deficit.
The plants with overexpressed CsPLDα exhibited lower water loss, higher leaf relative water content, and heavier fresh and dry matter accumulation than the WT. We proposed that CsPLDα was involved in the ABA-dependent pathway in mediating the stomatal closure and preventing the elevation of intracellular solute potential.
KeywordsCsPLDα Transgenic tobacco Phosphatidic acid Drought stress Abscisic acid Stomatal closure Lipid peroxidation
Liquid chromatography mass spectrometer
Nicotinamide adenine dinucleotide phosphate [H]
Quantitative real-time polymerase chain reaction
Reactive oxygen species
Relative water content
Environmental stresses trigger a wide variety of plant responses, and drought stress is one of the most adverse factors to plant growth and productivity [1, 2]. Drought stress can increase reactive oxygen species (ROS) generation, increase pyruvic acid content, decrease ascorbic acid content, induce lipid peroxidation injury, and cause irreversible damage, which leads to death . Plants form a complex regulatory mechanism to adapt or resist water deficit during prolonged evolution. Before drought occurs, plants accelerate tissue maturation to effectively avoid damage. Plants develop a strong root system by closing stomata to reduce water loss . External stimuli can be identified by plant cell membranes or osmosensors, leading to the production of secondary messengers that can be transported within the cell . The secondary messengers can adjust the phosphorylation status of downstream proteins by regulating the activity of intracellular protein kinase, invoking the activity of transcription factors, mediating the expression of target genes in the nucleus . These processes affect the plant morphogenesis, transformation of carbon metabolic pathways, hormone synthesis, ROS balance, and osmolyte accumulation, eventually enhancing plant resistance to stress [7, 8]. When plants sense a drought stress signal, intracellular signaling molecules, such as secondary messengers, are transmitted inside the cell. The signal transduction activates the abscisic acid (ABA)-dependent/independent pathways, which allow water deficit resistance . At least three ABA-dependent pathways exist in plants, and the transcription factors that regulate these pathways include myeloblastosis, NAC (NAM/ATAF1/2/CUC2), and others [10, 11]. ABA-independent pathway-related genes mainly include those that possess dehydrating response components in their promoter regions and transcription factors that combine with dehydration-responsive element/C-repeat components . Therefore, differences in ABA accumulation in plants induce different pathways in response to water deficit. Most of the genes related to ABA synthesis and degradation pathways including ZEP/ABA1, NCED, SDR/ABA2, LOS5/ABA3, AAO3/ABA5, CYP707A, and AOG, have been cloned and studied [13, 14, 15]. One study suggested that the SDR gene is not affected by the drought stress signal . Hence, ABA plays a vital role in the water deficit response mechanism of plants. ABA participates in the plant response to drought stress by controlling the opening and closing of plant stomata [17, 18]. The stress hormone ABA and elevated CO2 levels activate complex signaling pathways that are mediated by kinases/phosphatases, secondary messengers, and ion channel regulation in guard cells .
Some transcription factor genes respond to drought stress signal, but mainly two kinds of protein participate in the water deficit response: the regulatory and function proteins. The regulatory proteins include protein kinases, transcription factors, and phospholipases. These proteins are involved in the signal transduction of drought stress mainly by adjusting other signaling molecules. The function proteins include the LEA-like protein, molecular chaperones, osmolyte synthetases, transporters, and ROS detoxification protein enzymes. They are directly involved in the drought stress response and repair process . Among these proteins, the plant phospholipase D (PLD) family and the phosphatidic acid (PA) they produce function in drought stress responses [20, 21, 22, 23]. PLD exhibits the dual function of membrane degradation and signal transduction . As a lipid-hydrolyzing enzyme, PLD hydrolyzes membrane phospholipids to produce PA and exhibits increased activity under dehydration and hyperosmotic conditions [24, 25]. The produced PA acts as a secondary messenger, amplifying the signal to possibly mitigate stress injury; thus, it mainly functions in stress injury rather than in membrane degradation in some cases . The detrimental effects of drought stress are prevented by minimizing cuticular water loss and maximizing water uptake . PLDα1 mediates ABA regulation, which controls stomatal closure and decreases transpirational water loss in response to water deficits [28, 29]. Arabidopsis with abrogated PLDα1 is insensitive to ABA-mediated stomatal closure and exhibits more water loss than that of the wild type (WT) , whereas Arabidopsis with overexpressed PLDα1 loses less water than the WT . On one hand, PLDα1-produced PA binds to ABI1 protein phosphatase 2C, and this interaction may tether ABI1 to the plasma membrane, impede its negative function on ABA response, and enhance ABA-promoted stomatal closure [28, 29]. On the other hand, PLDα1 interacts with the Gα subunit of heterotrimeric G protein to mediate stomatal opening inhibition by ABA [29, 31]. PA also binds to NADPH oxidase and stimulates its activity to promote ROS or NO production in ABA-mediated stomatal closure . Additional studies are needed in order to further confirm these results.
Our previous study showed that the cucumber phospholipase D alpha gene (CsPLDα) was involved in the response to hyperosmotic stress, and the overexpression of CsPLDα in tobacco could enhance the tolerance to high salinity, polyethylene glycol and ABA treatments, which was proved in seed germination and seedling condition . Furthermore, the CsPLDα-produced PA participated in the salt response by congesting osmolytes, balancing Na+–K+ ratio and eliminating the accumulation of ROS indirectly . In the current study, we found that CsPLDα-produced PA could do more work in drought stress response, especially in promoting ABA-mediated stomatal closure, balancing osmolytes, and stabilizing the membrane system, which could keep more water in plants. Therefore, these findings suggested that CsPLDα contributed significantly to drought stress in plants.
NtPLDα1 and NtNAC072 expression under drought stress
CsPLDα mediated the plant response to ABA synthesis and metabolism, stomatal closure, and water loss
FW and DW under drought stress
EL and MDA content under drought stress
Soluble sugar, proline, and soluble protein contents under drought stress
CsPLDα expression improved tolerance to drought in tobacco
This study showed that the CsPLDα overexpression exerted positive effects to the plant response to water deficit. When external water is lacking, the water inside the plant cell leaks to the outside of the cell, decreasing cell turgor pressure and changing the osmotic potential across the cell membrane. According to the osmosensor hypothesis, a change in osmotic potential leads to a structural change of the kind of protein in cell membranes called osmosensors; reduced hyperosmolality-induced [Ca2+]i increase 1 (OSCA1) may be an osmosensor in Arabidopsis and is induced by a stimulus in plants . The main osmosensor in plants is a two-component system that includes histidine protein kinase and response regulator protein, and this system plays an important role in the rapid acceptance and transduction to osmotic signal [36, 37]. Aquaporins, one of the most important types of transmembrane osmosensors, can change the structure across the membrane by sensing differences in extracellular water potential and transfer the water potential signal into other intracellular signaling molecules . Through this process, the plant cell receives and conducts the drought signal through various pathways. The regulation of cell membrane lipid modification by phospholipase is one of the important pathways. Such modification can induce cells to produce different kinds of signaling molecules, such as PA, diacylglycerol (DAG), DAG-pyrophosphates, lysophosphatide, free fatty acids, and phosphatidylinositol [39, 40, 41].
PA plays an important role in the phospholipid signaling pathway and is mainly produced by phospholipase C and PLD . Water deficit can quickly activate PLD [43, 44], and the activated PLD is involved in ABA-mediated stomatal closure, which helps reduce transpiration and prevent water loss, meanwhile, maintains cell turgor pressure to against drought stress [28, 29]. PLDα1 and PA regulate ABA-induced stomatal closure by depolymerizing microtubules . PLDδ is also involved in ABA-induced stomatal closure. PLDδ mRNA levels increase in response to severe dehydration , and its expression is upregulated by ABA in guard cells . The interaction between PLDδ and glyceraldehyde-3-phosphate dehydrogenases in mediating plant response to ABA and water deficits has been well identified . Zhang et al.  suggested that PLDα1 and PLDδ are involved in the same signaling pathway activated by ABA. However, Uraji and Murata  proposed that PLDα1 and PLDδ demonstrate a cooperative function in ABA-induced stomatal closure. Moreover, PLDα1-deficient plants display delayed ABA-promoted leaf senescence . In the current study, CsPLDα was overexpressed in the tobacco plants, and the synthesis and metabolism of ABA in both transgenic and WT tobacco under drought treatment were monitored. First, the expression of endogenous NtPLDα1 exhibited no difference between the WT and transgenic plants, and the expression of the drought-induced marker gene NtNAC072 was improved in the transgenic plants under drought stress (Fig. 1). This finding suggested that the CsPLDα overexpression could enhance the sensitivity of plants to drought stress. The expression of the ABA-related genes was affected, the most important one of which was NtNCED1, which is a rate-limiting enzyme in the ABA synthesis pathway. The NtNCED1 expression was up-regulated by drought stress and was higher in the transgenic plants than in the WT. The expression of NtAOG, which mediates ABA degradation, was lower in the transgenic plants than in the WT. Thus, the ABA content significantly accumulated in the transgenic plants (Fig. 2). A gene in the ABA synthetic pathway, NtSDR, was not affected by the drought signal (Fig. 2c), and the previous research has supported the same result . Leaf stomatal closure extent was also observed. The stomatal aperture width of the transgenic plants was less than that of the WT (Fig. 3b). Thus, the increased stomatal closure in the transgenic plants reduced the water loss in comparison with the WT and kept relatively high level water content . The data on leaf water loss after a short water deficit (Fig. 3c) and leaf relative water content (Fig. 8b) after prolonged water deficit both supported this finding. These results indicated that CsPLDα was involved in the ABA-promoted stomatal closure during water deficit, further reduced the transpiration, and therefore enhanced the water retention.
Second, PA not only mediated the ABA-promoted stomatal closure to resist drought stress. In Arabidopsis, PLD can be used as a positive component to promote proline synthesis . Proline is one of the most effective intracellular osmolytes; it can enhance water retention and maintain membrane structure stability and enzyme activity [52, 53]. Therefore, proline content can directly or indirectly reflect the strength of the resistance of plants to osmotic stress. Other osmolytes maintain osmotic balance, including soluble sugar [54, 55] and glycine betaine . Soluble sugar content is increased in enhanced ZmPLC1 expression in transgenic maize . In the current study, the contents of proline, soluble sugar, and soluble protein in both the transgenic plants and WT accumulated rapidly and enhance the resistance to drought stress. However, in prolonged drought stress, the osmolyte content in the transgenic plant was more than that in the WT (Fig. 6); hence, the solute potential in the transgenic plant was less than that in the WT (Fig. 8a). These results showed that the CsPLDα-driven PA could mediate the osmolyte synthesis to maintain the osmotic balance and alleviate the stress damage.
MDA is commonly used to indicate lipid peroxidation and increases in response to short-term drought stress [58, 59]. EL is equally useful in stress . PLD is also involved in membrane lipid remodeling and rearrangement, which contribute to the synthesis of other kinds of lipids under stress, especially in phosphorus deficit, to maintain the balance of membrane system structure and cell function [61, 62, 63, 64]. In the current study, the MDA content and EL in the leaves of the transgenic plants were significantly lower than those of the WT (Fig. 5). This finding indicated that the overexpressed CsPLDα could reduce the damage of membrane lipid peroxidation, maintain the membrane system structure stability and function, and decrease the membrane ion leakage during water deficit.
Both overexpressed CsPLDα and it produced-PA are involved in controlling water loss by regulating more stomatal closure and the homeostasis of solute potential, therefore, CsPLDα is expressed dominantly in vigorously growing tobacco cells under drought stress, both in leaves and roots, and its overexpression plants can improve the tolerance to water deficit. These also indicated the role of CsPLDα in the regulating mechanism of plant response to drought stress is extremely complex, thus we still need further study to verify the interaction between all these different functions.
Plant materials, growth condition, and drought stress treatment
Tobacco (Nicotiana tabacum cv. NC89) was used for the assays as the wild type, and the transgenic lines ‘T1–68’ and ‘T1–71’ resulting from self-fertilization were produced during a previous research . Based on the previous study [33, 34], the WT seeds were sown 2 days earlier, and all seeds were sterilized and sown into plastic plots. Seedlings were grown in a controlled environment with an air temperature of 28 °C (day)/ 23 °C (night), a light intensity of 140 μmol m− 2 s− 1, and a relative humidity of 60%. The four-week seedlings were transferred into a plastic plug filled with nursery substrate, and grew in glass greenhouse at Shandong Agricultural University. The plants were watered with a Hoagland complete nutrient solution just like describing at  before treatment and refreshment. The experiment was conducted under natural conditions with an air temperature of 25–30 °C during the day and 18–25 °C at night. After 2 weeks, the young tobacco seedlings were used as experimental materials for the treatments. The drought stress treatment was performed according to the method described by Hong et al. . The plugs were arranged in a completely randomized block design with 15 replicates per treatment. The tobacco plants were watered with the Hoagland nutrient solution the day prior to the treatment (0 day), and the watering was stopped in the succeeding 30 days. All lines were refreshed for 2 days. The samples were collected after 0, 8, and 16 days of treatments, and quantitative real-time PCR (RT-PCR) were performed. The H2O2 and ABA contents; plant fresh weight (FW) and dry weight (DW); electrolytic leakage (EL); malondialdehyde (MDA), proline, soluble sugar, and soluble protein contents; relative water content (RWC); and solute potential were measured in the leaves or roots.
Stomatal closure and water loss determination
To determine the percentage of closed stomata and stomatal aperture in response to dehydration, the leaves of the tobacco seedlings prior to the treatment were detached and exposed to an illuminated incubator with an air temperature of 23 °C and a cool, white light intensity of 125 μmol m− 2 s− 1. At 0, 10, 20 and 30 min after detachment, the percentage of closed stomata and stomata aperture were determined using a scanning electron microscope (Japanese Electronics Co., Ltd.) and analyzed using the Image-Pro software. Each treatment in each genotype used 30 stomata. Water loss was determined after 0, 0.5, 1, 3, 5, and 8 h.
Total RNA was extracted from the leaves by using the TRIzol method in accordance to the manufacturer’s instructions (CWBio), and the reverse transcription reaction was done by using the TransScript One-Step gDNA Removal and cDNA Synthesis SuperMix (TransGen). The primer sequences are shown in Additional file 1: Table S1. For the determination of drought stress-associated gene expression, the qRT-PCR was performed using the TransStart® Tip Green qPCR SuperMix, and detected by using an ABI 7500 RT-PCR instrument (Thermo Fisher Scientific, USA), as described in the literature . Each expression profile was independently verified in triplicate. Data were analyzed using the SDS 2.0 software (ABI), and the relative gene expression levels were calculated using the 2−△△Ct method .
Extraction and determination of ABA content
The ABA extraction and determination were performed in accordance with the method of Cheng et al.  with modifications. Fresh samples (0.5 g) were homogenized and extracted, then incubated at 4 °C. Phosphate buffer was added to the extract, and 1 ng of ABA (SA8750, Beijing Solarbio Life Sciences, Beijing, China) was added to each sample as internal standard, in accordance with the method of Asami et al. . After distilling the acetone, lipids were removed by partitioning the aqueous concentrate with hexanes. Then the aqueous phase was adjusted to a pH 2.5 and extracted using ethyl acetate. The acidic fraction was dried and dissolved in methanol. The solution was subjected to HPLC on a μBondapak C18 (30 cm × 0.78 cm column; Waters, Milford, MA, USA). The ABA was collected from 10.0 min to 12.0 min. The fractions containing ABA were dried and methylated with diazomethane. The methylated ABA was used for the LC-MS analysis.
The LC-MS analysis was performed using a triple quadrupole liquid chromatography mass spectrometer (TSQ Quantum) with a DB-1 capillary column. The chromatographic conditions were as follows. Thermo Scientific Hypersil C18 column was used with mobile phases of methanol (B) and water (D). In the elution gradient, the mobile phase B was increased from 20 to 90% within 6 min, maintained at 90% for 2 min, and then decreased to 20% within 4 min. The velocity, column temperature, and sample quantity were 0.3 ml/min, 30 °C, and 5 μl, respectively. The mass spectrometry conditions were as follows. The negative ion mode was utilized for electrospray power. The spray voltage, gasification temperature, sheath pressure, auxiliary air pressure, ion transport tube temperature, collision gas, and scanning mode were 3.5 kV, 350 °C, 35 arb, 15 arb, 350 °C, 1.5 mTor, and SRM, respectively.
Determination of plant FW and DW
Under the stress conditions after 0, 8, and 16 days of treatment, three plants each from the WT, ‘T1–68’, and ‘T1–71’ were collected. The FWs of tobacco were measured immediately. The samples were dried at 80 °C for 48 h, and then the DWs were measured.
EL and lipid peroxidation analysis
The EL and MDA content were determined in accordance with the method of Wang et al.  with modifications. Leaves were sampled at 0, 8, and 16 days after treatment and washed with deionized water. The membrane ion leakage was expressed as the percentage of initial conductivity versus total conductivity. The MDA content was expressed as the nonspecific absorbance at 600 nm was subtracted from the absorbance at 532 nm, and the difference was used to calculate the amount of MDA by using an extinction coefficient of 155 mM− 1 cm− 1 .
Determination of proline, soluble sugar, and soluble protein contents
Proline extraction was used an improved sulfo-salicylic acid method which described by Xu et al. . The soluble sugar was determined by using sulfuric acid–anthrone colorimetry . The protein content determination was based on Bradford .
The FWs of the tobacco leaves were measured immediately after detachment from the seedlings. The turgor weight was determined after incubation in deionized water overnight, and the leaf samples were dried at 80 °C for 48 h to obtain the DW. The RWC was calculated based on the following equation: RWC (%) = (FW − DW) / (TW − DW) × 100 .
Measurement of solute potential
The solute potential (Ψs) was measured as described by Gaxiola et al. . The osmotic potential was determined using a vapor pressure osmometer (model 5520, Wescor, USA). Ψs = −moles of solute (R × K), where R = 0.008314 and K = 293 °.
Values were presented as means ± standard deviations of the triplicates. Statistical analyses were conducted using ANOVA with SAS (SAS Institute, Cary, NC, USA). Differences between treatments were separated by the least significant difference test at P < 0.05 and P < 0.01.
This work was supported by National Natural Sciences Foundations of China (NO.31372060, NO.30900983), Collaborative Innovation Center of Fruit & Vegetable Quality and Efficient Production in Shandong, the China Agriculture Research System (CARS-25-D) and Collaborative Innovation Center of Protected Vegetable Suround Bohai Gulf Region. This article is supported by the Foundation of Shandong Provincial Young Teachers’ Development Plan. All these funding play roles in the design of the study and collection, analysis, and in writing the manuscript.
Availability of data and materials
The datasets used and/or analysed during the current study available from the corresponding author on reasonable request.
Conceived and designed the research: FY, SL, and TJ. Performed the research and analyzed the data: SL and TJ. LL contributed to the qRT-PCR assay and participated in other assays. MH contributed to the acquisition of transgenic lines. XW, MW, QS made key comments on the design of the trial, the article writing, and the revisions. YL and BG contributed to the improvement of the method, which including plant growth condition, stomatal closure and ABA content determination. Wrote the first draft of the manuscript: TJ. Improved the first draft of the manuscript: FY and SL. All authors have read and approved this manuscript.
Ethics approval and consent to participate
The plant materials (including seeds) performed in the current study were complied with institutional, national and international guidelines. All of our researches were conducted in our laboratories and greenhouses at Shandong Agricultural University (Tai’an, China). No specific permits were required.
Consent for publication
The authors declare that they have no competing interests.
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