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First published online March 25, 2008; 10.1105/tpc.107.056390 The Plant Cell 20:803-816 (2008) © 2008 American Society of Plant Biologists
Phospholipase D
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ABSTRACT |
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 TOP ABSTRACT INTRODUCTIONRESULTSDISCUSSIONMETHODSREFERENCES |
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3 alter plant response to salinity and water deficit. PLD3 uses multiple phospholipids as substrates with distinguishable preferences, and alterations of PLD3 result in changes in PA level and membrane lipid composition. PLD3-KO plants display increased sensitivities to salinity and water deficiency and also tend to induce abscisic acid–responsive genes more readily than wild-type plants, whereas PLD3-OE plants have decreased sensitivities. In addition, PLD3-KO plants flower later than wild-type plants in slightly dry conditions, whereas PLD3-OE plants flower earlier. These data suggest that PLD3 positively mediates plant responses to hyperosmotic stresses and that increased PLD3 expression and associated lipid changes promote root growth, flowering, and stress avoidance. |
INTRODUCTION |
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TOPABSTRACTINTRODUCTIONRESULTSDISCUSSIONMETHODSREFERENCES |
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). Complex changes in gene expression, cellular metabolism, and growth patterns occur in plants in response to hyperosmotic stresses (Zhu, 2002; Bray, 2004). Several classes of regulatory components, including plant hormones, transcription factors, protein kinases, and Ca2+, have been identified as mediating plant responses to salinity and water deficit (Jonak et al., 2002; Zhu, 2002; Fujita et al., 2006). Despite great progress being made toward understanding the abiotic stress signaling pathways, little is known about the process by which hyperosmotic stress is sensed at cell membranes and transduced into physiological responses (Chinnusamy et al., 2004; Fujita et al., 2006).
Cell membranes play key roles in stress perception and signal transduction. Increasing results indicate that membrane lipids are rich sources for signaling messengers in plant responses to hyperosmotic stresses (Wang, 2004; Testerink and Munnik, 2005; Wang et al., 2006). In particular, phospholipase D (PLD), which hydrolyzes membrane lipids to generate phosphatidic acid (PA) and a free head group, is activated in Arabidopsis thaliana in response to various hyperosmotic stresses, such as high salinity (Testerink and Munnik 2005), dehydration (Katagiri et al., 2001), and freezing (Welti et al., 2002; Li et al., 2004), as well as abscisic acid (ABA), a phytohormone regulating plant water homeostasis (Zhang et al., 2004). In addition, PLD-produced PA increases rapidly in cell suspension cultures of tomato (Solanum lycopersicum) and alfalfa (Medicago sativa) subjected to salt stress and in dehydrated leaves of the resurrection plant (Craterostigma plantagineum) (Frank et al., 2000; Munnik et al., 2000). The changes in PLD activity, expression, and PA formation under these conditions imply a role for PLD in response to salinity and other hyperosmotic stresses. However, the physiological effects of PLD-mediated signaling and the identity of specific PLD(s) involved in plant responses to salinity and water deficiency remain to be determined.
Arabidopsis has 12 identified PLDs that are classified into six types, PLD (3), -β (2), -
(3), -
, -
, and -
(2) (Qin and Wang, 2002; Wang, 2005). Several PLDs have been implicated in specific physiological processes. PLD1 is the most abundant PLD in Arabidopsis tissues and is also more extensively characterized than other PLDs. PLD1 deficiency renders plants insensitive to ABA in the induction of stomatal closure (Zhang et al., 2004; Mishra et al., 2006). PLD1-derived PA binds to ABI1, a negative regulator of ABA signaling, to regulate water loss through stomata. PLD1 also interacts with the plant G protein through its DRY motif (Zhao and Wang, 2004). PLD is involved in freezing tolerance (Li et al., 2004) and dehydration-induced PA formation (Katagiri et al., 2001). PLD1 and -2 are involved in plant responses to phosphate deficiency (Cruz-Ramirez et al., 2006; Li et al., 2006), and PLD2 is also part of the auxin response (Li and Xue, 2007). The above distinct physiological effects resulting from the loss of one PLD indicate that individual PLDs have specific functions (Wang et al., 2006). However, except for PLD1, which has a role in the ABA regulation of stomatal movement and water loss, none of the characterized PLD mutants exhibit an overt phenotype under conditions of high salinity or drought.
The PLD group has three members; PLD1 and -2 are very similar, sharing 
93% similarity in deduced amino acid sequences, whereas PLD3 is more distantly related to the other PLDs, sharing 70% amino acid sequence similarity to each of the other two PLDs. The coding region of PLD3 contains three introns, whereas the coding regions of PLD1 and PLD2 are interrupted by two introns (Qin and Wang, 2002). This study was undertaken to characterize the biochemical properties and metabolic and physiological functions of PLD3. The results show that manipulations of PLD3 alter plant responses to hyperosmotic stress and indicate that PLD3 positively mediates plant responses to hyperosmotic stress.
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RESULTS |
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TOPABSTRACTINTRODUCTIONRESULTSDISCUSSIONMETHODSREFERENCES |
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31, but none for PLD3, indicating that the level of PLD3 expression is much lower than that of PLD1. This is supported by quantitative real-time PCR data showing that the average level of PLD3 expression in buds, flowers, siliques, stems, old leaves, and roots is 1000-fold lower than that of PLD1. Otherwise, the expression patterns in the different organs were similar between the two PLD genes (Figure 1A
). These results are consistent with the expression levels and patterns of PLD1 and PLD3 expression in different organs as determined by querying GENEVESTIGATOR (https://www.genevestigator.ethz.ch).
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3 encodes a functional PLD, the gene was tagged at the C terminus with hemagglutinin (HA) and expressed in Arabidopsis (Figure 1B). HA-tagged PLD3 was purified, and PLD activity was assayed at Ca2+ concentrations and conditions previously defined for PLD1, -β, -, and - (Pappan et al., 1998; Wang and Wang, 2001; Qin and Wang, 2002). PLD3 was active under PLD1 reaction conditions that included 50 mM Ca2+, SDS, and single-lipid-class vesicle (Figure 1C). PLD3 was inactive under PLDβ, -, or - conditions, which included phosphatidylinositol 4,5-bisphosphate (PIP2), phosphatidylethanolamine (PE), and micromolar levels or no Ca2+ in the reaction mixtures. PLD3 displayed low activity under PLD conditions that included micromolar levels of Ca2+ and oleic acid (Figure 1C). PLD3 hydrolyzed the common membrane phospholipids phosphatidylcholine (PC), PE, phosphatidylglycerol (PG), and phosphatidylserine (PS), having the highest activity toward PC and the lowest toward PS (Figure 1D). PLD3 had no activity toward phosphatidylinositol (PI) or PIP2 when assayed with single-lipid-class vesicles.
Manipulations of PLD3 Alter Plant Sensitivity to Salinity
To investigate the cellular functions of PLD3, a T-DNA insertion mutant of PLD3 was isolated. The PLD3 knockout plant (PLD3-KO), designated pld3-1, has a T-DNA insertion in the second exon, located 739 bp from the start codon (Figure 2A
). The homozygosity of the mutant was confirmed by PCR using PLD3-specific primers and a T-DNA left border primer (Figure 2B). The mutation resulted in loss of the expression of PLD3, as indicated by the absence of a detectable PLD3 transcript by RT-PCR. Thus, pld3-1 is a knockout mutant (Figure 2C). The mutant allele cosegregated with kanamycin resistance and susceptibility in a 3:1 ratio, suggesting a single T-DNA insertion in the genome. In addition, >30 independent transgenic Arabidopsis lines overexpressing HA-tagged PLD3 (PLD3-OE) under the control of the cauliflower mosaic virus 35S promoter were generated, and the expression of PLD3-HA in the plants was confirmed by immunoblotting using HA antibodies (Figure 1B). A number of independent lines of OE plants were tested for their stress responses, and their physiological effects were correlated with the level of overexpression. For further characterization, two or three representative independent transgenic lines were used in each experiment.
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3-1 plants displayed no significant morphological alterations under control growth conditions. No apparent differences in growth and development were observed when seeds of these plants were germinated under nitrogen or phosphorus deficiency, water lodging, or in response to the growth regulators 1-aminocyclopropane-1-carboxylic acid, indole acetic acid, or cytokinin. However, pld3-1 was more sensitive to salt stress than was the wild type, whereas PLD3-OE was less sensitive. In the absence of NaCl, nearly 100% of seeds of all genotypes germinated within 2 d (Figure 2D). In the presence of 150 mM NaCl, the germination of pld3-1 seeds was delayed, whereas that of PLD3-OE was enhanced compared with wild-type seeds in the early stage of germination (Figure 2E). The seedling size and root length of PLD3-OE were greater than those of the wild type, whereas those of pld3-1 were smaller (Figure 2G). When NaCl was increased to 200 mM, the germination of pld3-1 was much slower, whereas that of PLD3-OE was faster than that of the wild type (Figure 2F). Introducing native PLD3 into the pld3-1 mutant (PLD3 complementation) restored the mutant phenotype to wild-type plants (Figures 2D to 2F), confirming that the changes observed in the insertion mutant were caused by the loss of PLD3.
To further characterize the salt stress response, 4-d-old seedlings germinated under non-salt-stress conditions were transferred to MS agar plates containing 50 or 100 mM NaCl. Primary root growth was inhibited in pld3-1 plants, and the length was 50% that of wild-type plants (Figures 3A and 3B
). PLD3-altered plants also differed from wild-type plants in the number of lateral roots (Figure 3C). One week after transfer to MS plates containing 50 mM NaCl, pld3-1 seedlings had significantly fewer lateral roots per plant than PLD3-OE or wild-type plants, and PLD3-OE plants had significantly more lateral roots per plant than wild-type plants (Figure 3C). PLD3-OE and wild-type plants had similar primary root lengths at the early stages of salt stress (Figure 3B), but PLD3-OE rosettes grew better than wild-type rosettes under prolonged salt stress (Figure 3D). The pld3-1 phenotype was restored to wild type after genetic complementation with PLD3 (Figures 3A and 3B).
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3-altered plants were grown in the same pots with wild-type plants. pld3-1 plants were more susceptible to salt stress than PLD3-OE or wild-type plants. After 2 or 3 weeks of salt stress, pld3-1 plants became yellow and eventually died, whereas wild-type and OE plants survived and grew to maturation (Figure 3E). The rate of ion leakage, an indicator of membrane integrity, in pld3-1 plants was much higher than in wild-type and PLD3-OE plants (Figure 3F). Chlorophyll content was also significantly lower in pld3-1 than in wild-type plants (Figure 3G). These results suggest that PLD3 is required for normal growth in the presence of salt.
Alterations in PLD3 Expression Change Plant Development under Water Deficit
To determine whether the alteration was specific to salinity, pld3-1, PLD3-OE, and wild-type seedlings were tested for their responses to other hyperosmotic stresses. In the presence of 8% polyethylene glycol (PEG), the growth of pld3-1 seedlings was inhibited, whereas PLD3-OE seedlings grew better than wild-type seedlings (Figures 4A and 4B
). Compared with wild-type seedlings, pld3-1 seedlings had 80% of the biomass accumulation and 20% shorter primary roots, whereas PLD3-OE seedlings accumulated 25% more biomass and had longer primary roots and more lateral roots (Figures 4C to 4E). These results indicate that ablation of PLD3 decreases plant response to hyperosmotic stress, in addition to salt stress specifically.
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3 KO and OE was investigated in plants grown in soil with limited water supply. Water deficits were imposed on wild-type, pld3-1, and PLD3-OE plants at 25 to 30% of soil water capacity (soil saturated with water). Under water deficit, the relative water content of the leaves was 60% that of well-watered plants. Plants continued growing, but growth was slower than for plants grown under well-watered conditions. When water deficiency was chronic, PLD3-OE plants flowered earlier and pld3-1 plants flowered later than wild-type plants (Figures 5A, 5C, and 5D). On average, OE plants bolted and flowered 9 d earlier than wild-type plants, but pld3-1 plants flowered 6 days later than wild-type plants. At the time of flowering, OE plants had four and eight fewer rosette leaves than wild-type and pld3-1 plants, respectively (Figure 5D
). The flowering time was also affected by the level of PLD3 protein; the OE line with a higher level of PLD3 flowered earlier than did plants with a lower level of PLD3 (Figures 1B and 5B). The OE plants set seeds earlier and had more siliques than wild-type plants and plants containing the empty vector at the flowering stage (Figure 5E). However, under well-watered growth conditions, wild-type, pld3-1, and PLD3-OE plants displayed no differences in flowering time or in the number of rosette leaves or siliques.
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; Mathieu et al., 2007). Increases in the expression of FT promote flowering. Thus, we measured the expression patterns of FT and its paralogues, BROTHER OF FT AND TFL1 (BFT) and TWIN SISTER OF FT (TSF), by real-time PCR. Under well-watered conditions, the expression levels of FT and BFT were not different among 3-week-old PLD3-altered and wild-type plants, but levels of TSF were lower in pld3-1 than in wild-type plants. Under water deficit conditions, the FT expression level was lower in pld3-1 plants, whereas the expression levels of BFT and TSF were higher in OE plants than in wild-type and pld3-1 plants at the inflorescence stage (Figures 5F to 5H). The trend of changes in the expression of flowering timing markers is consistent with the different flowering times resulting from PLD3 alterations.
Changes in ABA Content and ABA Response under Osmotic Stress
The transition from vegetative to reproductive development is controlled by multiple environmental and endogenous factors. The hormone ABA regulates stress responses, flowering, seed germination, and development. ABA is induced by drought stress and inhibits plant flowering (Bezerra et al., 2004; Razem et al., 2006). To investigate whether alterations of PLD3 changed ABA level and ABA response, ABA content was measured in pld3-1, OE, and wild-type plants under control and drought conditions (Figure 6A
). Under control growth conditions, the ABA content of OE plants was 5% higher than that of wild-type plants, whereas the ABA content of pld3-1 plants tended to be lower than that of wild-type plants, although the difference was not significant. When water was withheld, increases in ABA occurred in all three genotypes. However, compared with day 0 of the same genotype, the significant increase occurred at 2 d earlier in OE plants than in pld3-1 and wild-type plants (Figure 6A, top panel). At 8 d without watering, all genotypes had similar levels of ABA. These results indicate that altered expression of PLD3 has a small, yet significant, effect on the basal level of ABA and that plants with ablation or elevation of PLD3 are still capable of the drought-induced accumulation of ABA.
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3-1, and OE plants under control growth conditions. However, RD29B expression in pld3-1 was increased greatly at day 6 without water, 2 d sooner than the expression increased in wild-type plants (Figure 6A, middle panel). In OE plants, increases in RD29B expression also occurred, but the magnitude was much smaller than in wild-type and pld3-1 plants, even after 8 d without water. Likewise, the expression of RAB18, another ABA-inducible gene, also exhibited an earlier and larger increase in pld3-1 than in wild-type plants, whereas that of OE plants was less induced by water deficit (Figure 6A, bottom panel).
When seedlings were grown on MS medium supplemented with ABA, the growth of pld3-1 seedlings was more inhibited than that of wild-type seedlings, whereas that of OE seedlings was less inhibited (Figure 6B). The biomass accumulation of pld3-1 was only 46% of wild-type levels, whereas that of OE was 145% of wild-type levels after 30 d of growth on MS medium containing 5 µM ABA (Figure 6C). Without ABA, all three genotypes accumulated similar amounts of biomass (Figure 4C). PLD1 has been shown to be involved in the promotion of stomatal closure by ABA (Zhang et al., 2004; Mishra et al., 2006). KO of PLD1 impeded stomatal closure and increased leaf water loss, but the water loss from detached leaves was not significantly different among PLD3-KO, PLD3-OE, and wild-type plants (Figure 6D). These results suggest that unlike PLD1, PLD3 is not involved in the ABA regulation of stomatal movement and transpirational water loss.
Effects of PLD3 on PA Content and Lipid Composition
PLD3 hydrolyzed various membrane phospholipids in vitro to produce PA (Figure 1D). To determine the effect of PLD3 on lipid composition, we quantitatively profiled glycerophospholipids and galactolipids of wild-type, pld3-1, and OE plants. Under control growth conditions, the levels of PC, PE, PG, PS, monogalactosyldiacylglycerol (MGDG), and digalactosyldiacylglycerol (DGDG) were similar in pld3-1 and wild-type plants. PA content in pld3-1 plants was 80% that of wild-type plants (Figure 7A
), indicating that PLD3 contributes to the production of basal PA.
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3-1 plants had higher levels of nearly all lipids, except PA, which was 60% of the wild-type level (Figure 7A). By comparison, under the same water stress conditions, the effect of PLD1 KO on lipid change was smaller than that of PLD3 KO. The level of PG was higher and that of PA was lower in PLD1-KO than in wild-type plants (Figure 7A).
The difference in PA among pld3-1, OE, and wild-type plants was due primarily to differences in levels of 34-carbon PA species, which contain 18- and 16-carbon fatty acids (Devaiah et al., 2006) (Figure 7B). In terms of potential substrate lipids, pld3-1 had higher levels of both 34- and 36-carbon PCs as well as higher levels of PG and PI, although PI was not a substrate in vitro (Figure 1D). 34:6 MGDG and 36:6 DGDG were also higher in pld3-1, and 34:6 PA, which is likely to be formed by the hydrolysis of 34:6 MGDG, was lower in pld3-1 plants. These results indicate that PLD3 is involved in the drought-induced loss of glycerol polar lipids and changes in membrane lipid composition.
Changes in the Levels of TOR and AGC2.1 Expression
PLD-derived PA has been shown to activate mammalian target of rapamycin (mTOR) signaling, which regulates protein synthesis, cell growth, and stress responses (Fang et al., 2001). TOR plays important roles in cell growth and embryonic development in Arabidopsis, as well as in hyperosmotic stress (Menand et al., 2002; Mahfouz et al., 2006). Our results show that alterations of PLD3 changed PA level, osmotic tolerance, growth, and development under salt and water deficit stresses. These observations led to testing of whether alterations of PLD3 affected the TOR signaling pathway in the hyperosmotic response. The transcript level of TOR in PLD3-altered plants was assessed under both salt stress and water deficiency conditions by real-time PCR. The level of TOR expression was lower in pld3-1 plants and higher in OE plants than in wild-type plants under both conditions (Figure 8A
). We also monitored the expression of AGC2.1 kinase, whose activity was shown to be regulated by PA to promote root hair growth in Arabidopsis (Anthony et al., 2004). The transcript level of AGC2.1 kinase was significantly lower in pld3-1 than in wild-type and OE plants under salt stress, but there was no difference in AGC2.1 expression between PLD3-altered and wild-type plants under water deficient conditions (Figure 8A). These results suggest that alterations of PLD3 affect the expression of AGC2.1 and TOR differently.
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). To investigate whether the level of phosphorylated S6K was changed in PLD3-altered plants, the proteins extracted from KO, OE, and wild-type plants were immunoblotted with an anti-phospho-p70 S6K antibody. Under control growth conditions, the level of phosphorylated S6K was not significantly different among KO, OE, and wild-type plants. However, under salt and water deficit conditions, the level of phosphorylated S6K was lower in pld3-1 plants than in wild-type plants (Figure 8B). OE and wild-type plants had similar levels of phosphorylated S6K under the 100 mM NaCl condition, and OE plants had a higher level than wild-type plants under the water deficit condition (8% PEG) (Figure 8B). Thus, the level of phosphorylated S6K is correlated with hyperosmotic tolerance. |
DISCUSSION |
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TOPABSTRACTINTRODUCTIONRESULTSDISCUSSIONMETHODSREFERENCES |
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3 alters the plant response to hyperosmotic conditions. PLD3 is most active under the conditions defined for PLD1 and has the highest activity toward PC among the various lipids tested. By comparison, in vitro assays show that PLD1 uses PC and PE almost equally well and has almost no activity toward PS (Pappan et al., 1998). In addition, the expression of PLD3 was much lower than that of PLD1; the difference in young leaves was 5000-fold. This observation is consistent with previous reports that PLD1-KO leaves have almost undetectable PLD activity under the PLD1 assay condition (Zhang et al., 2004). These results indicate that PLD3 and PLD1 have overlapping and yet distinguishable biochemical and regulatory properties and that activation of these PLDs may result in distinguishable hydrolysis of membrane lipids and changes in lipid composition under stress.
To gain insights into how PLD3 affects the plant response to osmotic stress, this study investigated the effect of PLD3 KO and OE on lipid composition, ABA responses, and cellular components involved in growth regulation and flowering time. Without applied stress, KO of PLD3 or -1 did not cause apparent changes in membrane glycerolipid composition, except that PA levels in the KO mutants were lower than those in wild-type plants. Under prolonged mild drought, PLD3-KO plants underwent less alteration of lipid composition than wild-type or PLD1-KO plants, meaning that under drought conditions, PLD3-KO plants were considerably different in lipid composition than wild-type or PLD1-KO plants. The greater effect on drought-altered lipid profiles of PLD3 ablation compared with PLD1 ablation was unexpected, given the finding that the expression and in vitro activity levels of PLD3 in leaves were much lower than those of PLD1. Indeed, ablation of PLD3 reduced the drought-induced decreases of almost all polar lipids, including PC, PE, PG, PS, PI, and DGDG (Figure 7), despite the fact that DGDG and PI are not substrates of PLD3. These results suggest that PLD3 promotes decreases in glycerolipids under water deficit but that much of the lipid loss in PLD3-KO plants does not result directly from PLD3-catalyzed hydrolysis. The notion that PLD3 acts in a regulatory role is consistent with the finding that, during drought, PLD3-OE and wild-type plants had similar levels of phospholipids and galactolipids, except for a lower level of PE. The specific effect of PLD3 on other lipolytic enzymes remains to be determined.
The results suggest that PLD3 plays a negative role in the plant response to ABA. However, KO and OE of PLD3 had no significant impact on leaf water loss (Figure 6D), suggesting that PLD3 is not involved in the ABA regulation of stomatal closure. By comparison, PLD1 has been shown to play a positive role in mediating the ABA promotion of stomatal closure and decreases in transpirational water loss (Zhang et al., 2004; Mishra et al., 2006). KO of PLD1 increases leaf water loss, whereas OE decreases the loss (Sang et al., 2001; Zhang et al., 2004). On the other hand, KO of PLD1 did not affect the ABA inhibition of seedling growth. The distinctively different effects of the two PLDs suggest that PLD1 and -3 enhance the plant osmotic stress response through different mechanisms. It might be possible that PLD1 mediates the ABA effect on stomatal movement to reduce water loss, whereas PLD3 promotes root growth in response to osmotic stress. Under hyperosmotic stress, PLD3-KO plants have shorter and fewer roots, whereas PLD3-OE plants have longer and more roots. A robust root system enables plants to maintain water status, thus delaying ABA-responsive gene expression. Thus, PLD3 may not be directly involved in the ABA signaling pathway and instead may be more involved in promoting growth under osmotic stress.
To test the effect of PLD3 KO and OE on plant growth, changes in TOR and S6K were monitored in PLD3-altered plants under hyperosmotic stresses. The TOR pathway is involved in the hyperosmotic stress response in animals (Fang et al., 2001) and also in plants (Mahfouz et al., 2006). Mammalian PLD1-derived PA was found to be a mediator in mTOR signaling. Our data show that pld3-1 plants had less PA than wild-type and OE plants under osmotic stress. Alterations of PLD3 led to a change in TOR transcription levels under both salt and water deficient conditions. In addition, the level of phosphorylated S6K was lower in pld3-1 plants than in wild-type plants under salt and water deficit conditions. TOR regulated cellular activities by activating downstream kinases, and S6K was one of the well-characterized TOR targets. PA activated mTOR, that stimulated S6K through phosphorylation. S6K has been implicated in the hyperosmotic stress response in Arabidopsis (Mahfouz et al., 2006). These results raise the possibility that PLD3 may be involved in the TOR signaling pathway in the stress response. One promising future direction is to determine whether PA interacts directly with TOR or S6K to regulate the hyperosmotic responses.
The altered stress responses exhibited in PLD3-OE and PLD3-KO plants may be caused by alterations in lipid metabolism and/or signal transduction. Changes in membrane lipid composition can result in changes of the localization and activities of signaling messengers associated with membranes. Specifically, the effect of PLD3 OE and KO on flowering time raises an interesting question: Do PLD3-mediated changes under water deficit affect key cellular components that control flowering time and the life cycle? Early flowering allows plants to accelerate their life cycle, an important mechanism by which plants escape stress. Arabidopsis flowering is controlled by environmental and endogenous signals (Corbesier et al., 2007). A key integrator of the signal input is FT, which encodes a small protein. In the current model, FT functions as a mobile signal moving from leaves to the shoot apex, where it interacts with the basic domain/Leu zipper transcription factor FLOWERING LOCUS D to activate the transcription of the floral meristem identity gene APETALA1. Interestingly, FT and its paralogues TSF and BFT contain a lipid binding domain with similarity to RAF kinase inhibitors that bind the membrane lipid PE (Mathieu et al., 2007). Under drought conditions, the level of PE in pld3-1 was higher, whereas that in OE was lower, than the wild-type level. Although increases in PC and PG and a decrease in PA occurred in pld3-1, PE was the only altered lipid class in OE plants. By contrast, KO of PLD1 did not change the PE level compared with the wild type (Figure 7). This raises the possibility that the altered PE levels may be responsible for the alteration of flowering time in PLD3-altered plants under water-deficient conditions. One scenario could be that the PE–FT interaction might tether the protein to membranes and attenuate its flowering-promoting functions. However, it is unknown whether FT or its paralogues actually interact with PE or other membrane lipids, such as PA.
Collectively, these results indicate that PLD3 plays a role in modulating plant growth and development under hyperosmotic stresses. The data suggest possible connections between membrane lipid–based signaling and some of the key regulators in flowering promotion and the hyperosmotic response. Further studies on the potential interactions of PLD and lipids with regulators such as ABA, FT, and TOR will help us better understand the mechanisms by which plants respond to salinity and water deficiency.
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METHODS |
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TOPABSTRACTINTRODUCTIONRESULTSDISCUSSIONMETHODSREFERENCES |
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3, designated pld3-1, was identified from the Salk Arabidopsis thaliana T-DNA knockout collection (SALK_130690), and seeds were obtained from the ABRC at Ohio State University. A PLD3 homozygous T-DNA insert mutant was isolated by PCR screening using the PLD3-pecific primers 5'-CTCGAGATGACGGACCAATTGCTGCTTCATCG-3' (forward primer) and 5'-ACGCCTAGAAGTAAGGATGATTGGAGGAAGA -3' (reverse primer) and the left border primer 5'-GCGTGGACCGCTTGCTGCAACT-3'. A pair of PLD3-specific primers were used in RT-PCR to confirm the PLD3 null mutant: 5'-ATGGTTAATGCAACGGCAGACGAG-3' (forward) and 5'-CCCGGTAAATCGTCATTTCGAGGA-3' (reverse). The PCR conditions were 95°C for 1 min for DNA melting, 40 cycles of 95°C for 30 s, annealing for 30 s (annealing temperature was based on the melting points of the specific primers), and 72°C for 30 s for DNA extension. Finally, the reaction was set at 72°C for 10 min. For complementation of the PLD3 knockout mutant, the native PLD3 gene, including its own promoter region, was amplified from 1.5 kb upstream of the start codon and 600 bp after the stop codon and then was cloned into the pEC291 vector. The primers for PLD3 complementation were 5'-CTGCAGGTAAGATTCACAAAATTGGTGTAATAC-3' (forward) and 5'-AAGCTTGAGTGAATATGGTCTATGGATATT-3' (reverse). The plasmid was transformed into pld3-1 plants with the flower-dipping method (Martinez-Trujillo et al., 2004). The transformants were selected from hygromycin plates and confirmed by PCR using the primers TeasyAsc5 (5'-ATGGCGCGCCATATGGTCGACCTGCAG-3') and TeasyAsc3 (5'-ATGGCGCGCCCGACGTCGCATGCTC-3'). PCR or RT-PCR products were visualized by staining with ethidium bromide on a 1% agarose gel after electrophoresis.
Plant Growth and Treatments
pld3-1, OE, wild-type, and pld3-1 complemented with PLD3 (PLD3 complementation) plants were grown in soil in growth chambers under 12-h-light/12-h-dark photoperiods (120 µmol·m–2·s–1) at 23/21°C and 50% humidity. For salt stress experiments, 3-week-old plants were treated with various concentrations of NaCl. Meanwhile, 4-d-old seedlings of pld3-1, OE, wild-type, and PLD3 complementation plants were transferred to MS (1x) agar plates containing 50 and 100 mM NaCl to test salt tolerance. For water stress experiments, 3-week-old plants (before inflorescence formation) were not watered for several days until soil water content was 25 to 30% of soil water capacity (soil saturated with water). The water-deficient condition was maintained by adding 50 mL of water to each pot (12 x 12 x 14 cm) every 4 d. Under this condition, the relative water content of leaves was 50%, whereas the relative water content for well-watered plants was 80%. For seed germination in response to osmotic stress or hormone treatment, seeds were germinated on MS (1x) agar plates supplemented with NaCl, PEG, or ABA. To minimize experimental variation, plants of similar size of different genotypes were grown in the same pots or on the same plates for stress treatments.
Expression, Purification, and PLD3 Activity Assay
The PLD3 gene was amplified from Arabidopsis genomic DNA using the PLD3 gene-specific primers 5'-CTCGAGATGACGGACCAATTGCTGCTTCATCG-3' (forward) and 5'-ACGCCTAGAAGTAAGGATGATTGGAGGAAGA 3' (reverse), introducing cloning sites of XhoI/StuI. The PLD3 sequence was fused with DNA encoding an HA tag and cloned into a binary pKYLX71 vector. HA-tagged PLD3 was expressed in Arabidopsis plants under the control of the 35S promoter. The C-terminally tagged PLD3-HA protein was purified from plant proteins by immunoaffinity column chromatography using HA antibodies conjugated to agarose beads. The purified protein was used for activity assays with dipalmitoylglycero-3-phospho-(methyl-3H) choline as a substrate under different conditions defined previously for other PLDs (Pappan et al., 1997; Wang and Wang, 2001; Qin and Wang, 2002). Briefly, PLD1 activity was assayed in the presence of 25 mM Ca2+, 100 mM MES, pH 6, 0.5 mM SDS, and 2 mM PC. PLDβ and - were assayed using 5 µM Ca2+, 80 mM KCl, 2 mM MgCl2, 100 mM MES, pH 7, and 0.4 mM lipid vesicle composed of PC:PE:PIP2 (0.2:3.5:0.3). The PLD reaction condition was 100 mM MES, pH 7, 2 mM MgCl2, 80 mM KCl, 100 µM CaCl2, 0.15 mM PC, and 0.6 mM oleate. PLD1 activity was measured in the presence of 100 mM Tris-HCl, pH 7, 80 mM KCl, and 0.4 mM lipid vesicle composed of PC:PE:PIP2 (0.2:3.5:0.3) (Qin and Wang, 2002). Hydrolysis of PC was quantified by measuring the release of [3H]choline by scintillation counting.
Real-Time PCR
Real-time PCR was performed as described by Li et al. (2006). Briefly, total RNA was extracted from leaves using the cetyl-trimethyl-ammonium bromide method. DNA was removed from RNA by digestion with RNase-free DNaseI. RNA was used as a template for reverse transcription to synthesize cDNA using the iScript kit (Bio-Rad). Quantitative real-time PCR was performed with the MyiQ sequence detection system (Bio-Rad) by monitoring the SYBR Green fluorescent labeling of double-stranded DNA synthesis. The efficiency of the cDNA synthesis was assessed by real-time PCR amplification of a control gene encoding UBQ10 (At4g05320), and the UBQ10 gene Ct value was 20 ± 0.5. Only cDNA preparations that yielded similar Ct values for the control genes were used for the determination of PLD gene expression. The level of PLD expression was normalized to that of UBQ10 by subtracting the Ct value of UBQ10 from the Ct value of PLD genes (Li et al., 2006). The primers for different genes were as follows: PLD3, 5'-ATGGTTAATGCAACGGCAGACGAG-3' (forward) and 5'-CCCGGTAAATCGTCATTTCGAGGA-3' (reverse); RD29B, 5'-ACAATCACTTGGCACCACCGTT-3' (forward) and 5'-AACTCACTTCCACCGGAATCCGAA-3' (reverse); RAB18, 5'-GCAGTCGCATTCGGTCGTTGTATT-3' (forward) and 5'-ACAACACACATCGCAGGACGTACA-3' (reverse); TOR, 5'-AGTTCGAAGGGCAAAGTACGACGA-3' (forward) and 5'-TACGCACGCTCATAGCTCTCCAAA-3' (reverse); AGC2.1, 5'-AGAAACGTCTCTTCCGCTTCACCA-3' (forward) and 5'-ACCTGAAGAATCTGACACGGCCAA-3' (reverse); FT, 5'-TCCCTGCTACAACTGGAACAACCT-3' (forward) and 5'-ACGATGAATTCCTGCAGTGGGACT-3' (reverse); BFT, 5'-ATTCAAACAGAGAGGGAGGCAAGC-3' (forward) and 5'-GCAGCAACAGGTTGAGAAAGACCA-3' (reverse); TSF, 5'-AAGACAAACGGTTTATGCACCGGG-3' (forward) and 5'-TTGAAGTAAGAGGCAGCCACAGGA-3' (reverse); and UBQ10, 5'-CACACTCCACTTGGTCTTGCGT-3' (forward) and 5'-TGGTCTTTCCGGTGAGAGTCTTCA-3' (reverse). The PCR conditions were as follows: 1 cycle of 95°C for 1 min; 40 cycles of 95°C for 30 s for DNA melting, 55°C for 30 s for DNA annealing, and 72°C for 30 s for DNA extension; and 72°C for 10 min for final extension of DNA.
Immunoblotting and Detection of Phosphorylated S6K
Total proteins were extracted from plants or seedlings grown in different conditions using buffer A (50 mM Tri-HCl, pH 7.5, 10 mM KCl, 1 mM EDTA, 2 mM DTT, and 0.5 mM phenylmethylsulfonyl fluoride). After centrifugation at 6000g for 10 min, the supernatant proteins were separated by 10% SDS-PAGE. After electrophoresis, proteins were transferred to a polyvinylidene difluoride membrane. The membrane was blotted with anti-HA antibody (1:1000) overnight, followed by incubation with a second antibody (1:5000) conjugated with alkaline phosphatase. The protein bands were visualized by alkaline phosphatase reaction. To detect phosphorylated S6K, proteins were transferred to nitrocellulose membranes and blotted with an anti-phospho-p70 S6K (Thr-389) antibody (Cell Signaling Technology), followed by a secondary antibody conjugated with horseradish peroxidase. The rabbit polyclonal antibodies were raised against human p70 S6K and have been shown to react with plant S6K proteins (Reyes de la Cruz et al., 2004). The membranes were preblotted with TBS/T containing 5% BSA and then were incubated with the first antibody (1:1000) in TBS/T buffer. After gentle agitation at room temperature for 1 h, the membranes were washed with TBS/T four times. A polyclonal anti-rabbit IgG antibody conjugated with horseradish peroxidase (1:10,000) was added and incubated for 1 h, followed by three washes with TBS/T and three washes with PBS. After incubation of LumiGLO substrate for 1 min, membranes were exposed to x-ray film.
Lipid Profiling and ABA Measurement
Lipid profiling was performed as described previously (Devaiah et al., 2006). Briefly, leaves were detached and immediately immersed in 3 mL of 75°C isopropanol with 0.01% butylated hydroxytoluene for 15 min, followed by the addition of 1.5 mL of chloroform and 0.6 mL of water. After shaking for 1 h, the extracting solvent was transferred to a clean tube. The leaves were reextracted with chloroform:methanol (2:1) five times with agitation for 30 min each, and the extracts were combined and then washed with 1 M KCl, followed by another wash with water. The solvent was evaporated with a stream of nitrogen. For each treatment, four leaf samples were extracted and analyzed separately. For ABA analysis, fresh leaves (100 mg) were ground in liquid nitrogen. Then, 0.5 mL of 1-propanol:H2O:HCl (2:1:0.002) was immediately added to the homogenate and mixed well. The homogenate was agitated at 4°C for 10 min, followed by the addition of 1 mL of dichloromethane and ABA internal standards. After vortexing and agitation at 4°C for 10 min, the mixtures were centrifuged at 11,300g for 1 min to separate the two phases. The lower phase was transferred to a 1.5-mL vial with a Teflon-lined screw cap. ABA was quantified by mass spectrometry as described by Pan et al. (2008).
Relative Water Content, Ion Leakage, and Chlorophyll
Leaves were detached and fresh weight (FW) was measured followed by incubation in clean water overnight to obtain turgor weight (TW). Leaves were then dried at 80°C for 48 h to measure dry weight (DW). The relative water content (RWC) was obtained based on the following equation: RWC (%) = (FW – DW)/(TW – DW) x 100. To measure ion leakage, leaves were detached and rinsed with distilled water and then were immersed in 15 mL of distilled water in glass tubes. After degassing under vacuum for 30 min to remove air bubbles on the leaf surface, samples were incubated with gentle agitation for 3 h (Fan et al., 1997). Initial conductivity was measured with a conductivity meter, and then the samples were boiled in a water bath for 20 min. Total conductivity was measured again after cooling to room temperature. Ion leakage was expressed as a percentage of the initial conductivity over total conductivity. For chlorophyll content measurement, chlorophyll was extracted from leaf discs placed in sealed vials with an appropriate volume of 100% methanol by shaking in the dark until the leaves became white. The chlorophyll content was obtained based on the absorbance of extracts at 650 and 665 nm (Crafts-Brandner et al., 1984).
Accession Numbers
Sequence data from this article can be found in the Arabidopsis Genome Initiative database under the following accession numbers: PLD3, At5g25370; RD29B, At5g52300; RAB18, At5g66400; TOR, At1g50030; AGC2.1, At3g25250; FT, At1g65480; BFT, At5g62040; TSF, At4g20370; UBQ10, At4g05320.
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Acknowledgments |
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Footnotes |
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www.plantcell.org/cgi/doi/10.1105/tpc.107.056390
Received October 19, 2007; Revision received January 30, 2008. accepted March 2, 2008.
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3 Is Involved in the Hyperosmotic Response in Arabidopsis
