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Keywords:

  • biomass;
  • plant growth;
  • nitrogen signaling;
  • phosphatidic acid;
  • phospholipase D

Summary

  1. Top of page
  2. Summary
  3. Introduction
  4. Results
  5. Discussion
  6. Experimental procedures
  7. Acknowledgements
  8. References

Activation of phospholipase D (PLD) produces phosphatidic acid (PA), a lipid messenger implicated in cell growth and proliferation, but direct evidence for PLD and PA promotion of growth at the organism level is lacking. Here we characterize a new PLD gene, PLDε, and show that it plays a role in promoting Arabidopsis growth. PLDε is mainly associated with the plasma membrane, and is the most permissive of all PLDs tested with respect to its activity requirements. Knockout (KO) of PLDε decreases root growth and biomass accumulation, whereas over-expression (OE) of PLDε enhances root growth and biomass accumulation. The level of PA was higher in OE plants, but lower in KO plants than in wild-type plants, and suppression of PLD-mediated PA formation by alcohol alleviated the growth-promoting effect of PLDε. OE and KO of PLDε had opposite effects on lateral root elongation in response to nitrogen. Increased expression of PLDε also promoted root hair elongation and primary root growth under severe nitrogen deprivation. The results suggest that PLDε and PA promote organism growth and play a role in nitrogen signaling. The lipid-signaling process may play a role in connecting membrane sensing of nutrient status to increased plant growth and biomass production.


Introduction

  1. Top of page
  2. Summary
  3. Introduction
  4. Results
  5. Discussion
  6. Experimental procedures
  7. Acknowledgements
  8. References

Membrane lipid hydrolysis by phospholipases (PL) such as PLD, PLC and PLA produces various classes of lipid mediators in growth response to nutrient and stress cues (Foster and Xu, 2003; Wang, 2004; Huang and Frohman, 2007). Activation of one or more of these reactions often constitutes an early, critical step in many signaling cascades. Phosphatidic acid (PA) has recently emerged as a new class of lipid messenger (Wang, 2004; Testerink and Munnik, 2005; Wang et al., 2006; Carman and Henry, 2007). PA has been shown to regulate proteins that are important in cellular regulation, including G proteins, protein kinases, phosphatases and transcriptional factors (Fang et al., 2001; Mishra et al., 2006; Carman and Henry, 2007; Zhao et al., 2007). PLDs are a major family of enzymes that generate signal molecule PA and play a critical role in regulating the location and timing of PA production (Wang et al., 2006). PLD and PA have been associated with mammalian cell growth, proliferation and survival signaling (Fang et al., 2001, 2003; Foster and Xu, 2003). In plants, PLD and PA have been implicated in promoting cell elongation in pollen and root hairs (Potocky et al., 2003; Anthony et al., 2004), as well as in plant growth in response to phosphorus deficiency and hyper-osmotic stress (Cruz-Ramirez et al., 2006; Li et al., 2006; Hong et al., 2008). However, the role of PLD and PA in promoting whole-organism growth under normal growth conditions has not been documented in any system.

PLDs are a heterogeneous family. The biochemical properties, domain structures and genomic organization of PLDs are more diverse in higher plants than in other organisms (Wang et al., 2006). The Arabidopsis genome contains 12 PLD genes (three α, two β, three γ, one δ, one ε and two ζ), ten of which have the Ca2+/phospholipid-binding C2 structural fold, whereas the two PLDζ proteins contain phosphoinositide-interacting pleckstrin homology (PH) and Phox homology (PX) domains (Qin and Wang, 2002). The presence of these regulatory motifs provides insights into the various modes of activation and functions of PLDs. Distinct biochemical properties and physiological functions have been reported for several PLDs. These include a role for PLDα1 in abscisic acid regulation of stomatal movement, for PLDα3 in hyper-osmotic tolerance, for PLDβ in defense response, for PLDδ in H2O2 response and freezing tolerance, and for PLDζs in root development in response to phosphate starvation and auxin (Zhang et al., 2003, 2004; Li et al., 2004, 2006; Cruz-Ramirez et al., 2006; Mishra et al., 2006; Li and Xue, 2007; Hong et al., 2008). The function of the other PLDs remains unknown.

PLDε encodes a protein that is distinctively different from the other 11 PLDs in Arabidopsis. It has the C2 structural fold, but contains no acidic residues involved in Ca2+ binding in the C2 domain (Qin and Wang, 2002). Phylogenetic analysis of the Arabidopsis and rice PLD families suggests that, of the C2 PLDs, PLDε is the one that is most closely related to the PX/PH PLDζs. Here, we characterize the properties and function of PLDε, and show that PLDε is more permissive than other characterized PLDs with regard to its activity requirements, and that genetic manipulation of PLDε alters Arabidopsis root growth and biomass accumulation. Further analysis of the PLD-altered plants indicated that PLDε and its derived PA are involved in N signaling, and that this lipid signaling may play a role in connecting membrane sensing of nutrient status to translational regulation of growth.

Results

  1. Top of page
  2. Summary
  3. Introduction
  4. Results
  5. Discussion
  6. Experimental procedures
  7. Acknowledgements
  8. References

PLDε is associated with membranes and is active under various reaction conditions

PLDε was expressed in all Arabidopsis tissues examined, and the PLDε mRNA level was highest in roots and low in leaves (Figure 1a). The relative level of PLDε expression in tissues was much lower than that of PLDα1 and the pattern of expression was also different from that of PLDα1 (Figure 1a). Expression data from Genevestigator (http://www.genevestigator.ethz.ch) also indicated that the level of PLDε expression was much lower than that of PLDα1 except in pollen, where expression of PLDα1 was low and that of PLDε was much higher than in other tissues. To determine whether PLDε encodes a functional enzyme, the protein was fused to a hemagglutinin (HA) tag at its C-terminus and expressed in Arabidopsis plants under the control of the CaMV 35S promoter. PLDε was then isolated from plants by immuno-affinity chromatography. The isolated PLDε–HA fusion protein was detected by the HA antibody, but not by an antibody raised against PLDα1 (Figure 1b). As a control, proteins from Arabidopsis leaves transformed with an empty HA vector were isolated using the same immuno-affinity procedure. No protein band was detected for the vector control using either HA or PLDα1 antibodies (Figure 1b). These results indicate that PLDε is purified without apparent contamination from the common PLDαs.

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Figure 1.  Expression and biochemical properties of PLDε. (a) Expression of PLDε and PLDα1 in Arabidopsis tissues as determined by real-time PCR. The expression levels were normalized in comparison to that of UBQ10. (b) Immunoblotting of PLDε isolated by immunoaffinity chromatography from plants transformed with 35S::PLDε–HA or an empty vector. Proteins were separated on 8% SDS–PAGE, blotted with anti-HA or anti-PLDα antibodies, and made visible by staining with alkaline phosphatase. (c) PLDε reaction conditions. The purified PLDε–HA was used for PLD activity assays under PLDα1, β, δ, and ζ reaction assay conditions. Values are means ± SD (= 3). (d) PLDε activity towards various phospholipids, NBD-PC, -PE, -PG or -PS, under PLDα1 reaction conditions. Values are means ± SD (= 3). Vector refers to a control in which proteins from Arabidopsis leaves transformed with an empty HA vector were isolated using the same immunoaffinity procedure used to isolate PLDε–HA.

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The purified PLDε was assayed under reaction conditions that were defined previously for PLDα1, β, δ and ζ, which display distinct requirements for Ca2+, polyphosphoinositides (PPI) and fatty acids, and substrate selectivity (Wang, 2004; Wang et al., 2006). PA production was proportional to the amount of enzyme and the reaction time. PLDε was active under the PLDα1 reaction conditions comprising 50 mm Ca2+, SDS and single-class lipid vesicles, but none of the other previously characterized enzymes PLDβ, γ, δ or ζ displayed activity under PLDα1 conditions (Qin and Wang, 2002) (Figure 1c). At micromolar concentrations of Ca2+, PLDε required oleic acid for activity, a requirement similar to that of PLDδ (Figure 1c). PLDε also displayed some activity under PLDβ and γ conditions, which included micromolar concentrations of Ca2+ and the lipids phosphatidylinositol 3,4-bisphosphate (PIP2) and phosphatidylethanolamine (PE) (Figure 1c). PLDε hydrolyzed the common membrane phospholipids phosphatidylcholine (PC), PE and phosphatidyl-glycerol (PG), and had low activity on phosphatidylserine (PS) when the enzyme was assayed with single-class lipid vesicles (Figure 1d). The results indicate that PLDε is active under a broad range of reaction conditions.

PLDε was present in the microsomal but not in the soluble fractions (Figure 2a). When the membrane fraction was separated into plasma and intracellular membrane fractions, most PLDε was associated with the plasma membrane (Figure 2a). To verify the subcellular localization, PLDε was fused with yellow fluorescence protein (YFP) at the C-terminus and transiently expressed in tobacco leaves. YFP alone was detected in the nucleus and cytoplasm as expected (Figure 2b, left) whereas PLDε–YFP fluorescence was detected on the plasma membrane (Figure 2b, right).

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Figure 2.  Subcellular localization of PLDε. (a) Subcellular fractionation of PLDε. S, soluble fraction; M, microsomal fraction; PM, plasma membrane; IM, intracellular membrane: 30 μg per lane were loaded for soluble proteins, and 5 μg per lane for membrane fractions. (b) Subcellular localization of PLDε–YFP using 35S_P:YFP:rbcS_T as a control (left) and 35S_P:AtPLDe:YFP:rbcS_T (right). The constructs were transiently expressed in tobacco leaves by infiltration. Scale bar = 50 μm.

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Over-expression of PLDε increases plant growth

PLDε-over-expressing (OE) plants, together with a homozygous knockout (KO) mutant pldε-1 (Figure 3a,b), were used to investigate the physiological functions of PLDε. More than 20 independent PLDε-OE lines were obtained, and production of PLDε–HA was confirmed by immunoblotting using HA antibody (Figure 3c). The KO mutant was isolated from the SALK_023603 line in the Arabidopsis Columbia ecotype. The mutant pldε-1 contained a T-DNA insertion in the second exon, 631 bp downstream of the start codon (Figure 3a). The mutation resulted in loss of PLDε expression, as indicated by the absence of detectable PLDε transcript (Figure 3b). The mutant allele co-segregated with kanamycin resistance and susceptibility in a 3:1 ratio. When seeds were germinated in hyper-osmotic growth media containing NaCl, sorbitol or 5–8% polyethylene glycol (PEG), pldε-1 seedlings grew more slowly and had shorter roots, whereas OE seedlings grew faster and had longer roots than wild-type (WT) seedlings (Figure 3d–g). The primary root lengths in KO plants were only 16, 70 and 60% of WT in 100 mm sorbitol, 5% PEG and 50 mm NaCl, respectively (Figure 3g). When seedlings were germinated in non-stress medium and then transferred to hyper-osmotic media, KO pldε-1 seedlings accumulated only 50 or 65% of the amount of dry matter accumulated by WT plants in the presence of 50 mm NaCl or 100 mm sorbitol, respectively (Figure 3h). Genetic complementation of the KO mutant with the PLDε gene restored the mutant phenotype to that of WT, confirming that the growth alterations result from loss of PLDε (Figure 3f–h).

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Figure 3.  PLDε and PA enhance growth under hyper-osmotic stress. (a) T-DNA insertion site in the PLDε gene. Boxes denote exons and lines introns. (b) Determination of PLDε transcripts in pldε-1 and WT plants by RT-PCR. UBQ10 was used as a control. WT, wild-type; pldε-1, PLDε knockout. (c) Immunoblotting of PLDε in 35S::PLDε–HA plants. Proteins extracted from leaves of plants transformed with the empty vector or 35S::PLDε–HA were separated by SDS–PAGE, followed by immunoblotting with HA antibodies. OE, PLDε over-expression. (d–f) Growth phenotypes of PLDε-altered seedlings under conditions of no NaCl (d), 50 mm NaCl (e) or 100 mm sorbitol (f). COM, PLDε knockout complementation. (g) Primary root growth of PLDε-altered and WT seedlings in response to osmotic conditions. PLDε-altered and WT seeds were germinated in MS (control) or MS containing 100 mm sorbitol, 5% PEG or 50 mm NaCl. The primary root length was measured at day 10 after sowing. Values are means ± SD (= 10) from one representative of three independent experiments. Each genotype contained 30 seedlings. (h) Effect of PLDε on biomass. Four-day-old seedlings were transferred to standard growth medium (control) or growth media containing 100 mm sorbitol or 50 mm NaCl without or with 0.15% 1-butanol or 2-butanol for 3 weeks, and were harvested for dry weight determination. Values are means ± SD (= 10) from one representative of three independent experiments. Each genotype was represented by 30 seedlings.

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In the process of analyzing PLDε-altered plants, we noted that PLDε-OE plants were larger than WT plants when grown on agar plates without stress treatment. The growth difference was also observed with PLDε-altered plants in soil (Figure 4a–c). The fresh and dry weights of rosettes of PLDε-OE were 192 and 212% those of WT, respectively, after five weeks of growth under well-fertilized conditions (Figure 4c). The increase in biomass resulted from increases in leaf size (Figure 4d) and leaf number (Figure 4e). PLDε-OE plants also had approximately 25% higher seed yield than WT (Figure 4f). Enhanced growth in PLDε-OE plants was also observed under less well-fertilized conditions (Figure 4c). The cell size of OE leaves was larger, whereas that of KO leaves was smaller than WT (Figure 4g). The magnitude of the increase in cell size was smaller than that in leaf area (Figure 4d), implying that cell number also increased in PLDε-OE plants. When OE, KO and WT plants were planted in the same tray, KO plants (Figure 4b, marked by arrows) were out-competed by WT and OE, and were much smaller in size than OE and WT plants.

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Figure 4.  Over-expressing PLDε enhances Arabidopsis growth in soil. (a) Rosettes of WT and PLDε-OE plants grown in soil. (b) Growth competition of PLDε-altered and WT plants. White arrows indicate KO plants. (c) Effect of fertilizer levels on rosette dry weight of 5-week-old plants. Values are means ± SD (= 30). (d,e) Leaf size and number for 5-week-old plants grown in well-fertilized soil. Values are means ± SD (= 40). (f) Seed yield. Values are means ± SD (= 10) from one of three independent experiments. (g) Cell sizes of expanded leaves from 5-week-old plants. Values are means ± SD (= 60). Asterisks indicate significant differences at < 0.05, compared to WT, based on Student’s t test.

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PLDε enhances root growth and biomass accumulation in response to nitrogen

Because of the growth differences, we investigated the effect of PLDε alterations on plant responses to macronutrients, nitrogen (N), phosphorus (P), potassium (K) and sulfur (S) using media with defined nutrient composition and concentrations. Seedlings were grown in agarose plates with the concentrations of the individual macronutrients reduced 10- and 100-fold from those of MS basal media. At 0.6 and 6 mm N (NO3:NH4+ = 2:1), the elongation of lateral roots was significantly higher in PLDε-OE than WT seedlings, but lower in PLDε-KO seedlings (Figure 5a,b). When N was maintained at 6 mm and P, S and K were reduced 100-fold to 13, 15 and 213 μm, respectively, WT and KO plants displayed similar rates of lateral root elongation. Under P- and K-limiting conditions, PLDε-OE roots elongated faster than those of WT, but the difference was not as great as that under N-deprived conditions (Figure 5a). When seedlings were grown on medium containing nitrate only, the elongation rate was also greater in OE and smaller in KO than in WT seedlings (Figure 5c). The difference was greater in seedlings grown in low levels of nitrate (0.6 and 6 mm) than in those grown in high levels of nitrate (60 mm) (Figure 5c). High levels of nitrate are known to have an inhibitory effect on lateral root development (Zhang et al., 1999; Walch-Liu et al., 2006). Genetic complementation of the KO mutant with the PLDε gene and its own promoter restored the mutant phenotype to that of WT, confirming that the growth alterations result from the loss of PLDε (Figure 5c).

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Figure 5.  Lateral root elongation of PLDε-altered seedlings responds differently to N, P, K or S deprivation. (a) Root elongation under various nutrient-deprivation conditions: N (0.6 mm), P (0.013 mm), S (0.015 mm) or K (0.213 mm). Seedlings were germinated and grown in agarose MS medium for 1 week, and then transferred to agarose MS media deprived of individual nutrients. Elongation of total lateral roots of individual plants was measured between 5 and 7 days after transfer. Except for N-deprivation conditions, all media contained 6.0 mm N. Values are means ± SD (= 30). (b) Seedling morphology of PLDε-OE, KO and WT plants. Four-day-old seedlings were transferred to MS salt containing 6 mm N for 12 days. (c) Lateral root elongation. Root length was measured 8 and 9 days after transferring 4-day-old seedlings to MS salt with 0.6, 6 and 60 mm nitrate. Values are means ± SD (= 60).

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PLDε-OE plants accumulated substantially more biomass, whereas KO had less biomass than WT when grown at various levels of N (0.6, 2, 6 or 60 mm) (Figure 6a). The mean dry weight of KO plants was 20% less than that of WT, whereas that of OE plants was approximately 20, 30 and 40% higher than in WT plants at N levels of 0.6, 2 and 6 mm, respectively. PLDε-OE plants grew more and longer lateral roots, whereas KO had fewer and shorter roots than WT (Figure 6b). The difference was greater in seedlings grown in low (0.6–6 mm) rather than high (60 mm) levels of nitrogen. At 6 mm N, the numbers and length of lateral roots of OE plants were twofold that of WT and KO plants (Figure 6b). The primary root length was no different in OE, WT and KO plants at 6 and 60 mm N, but roots of OE plants were about 20% longer than those of WT and KO plants under severely N-limited conditions (0.6 mm) (Figure 6b). When seeds were germinated directly under N-limited conditions (0.1 and 0.6 mm), the primary roots of PLDε-KO were shorter than those of WT, whereas those of PLDε-OE seedlings were longer than those of WT (Figure 7a,b). OE plants had more and KO plants had fewer lateral roots than WT (Figure 7c). These results suggest that basal PLDε gene function is required for maintaining normal primary and lateral root growth, particularly under N-limiting conditions. In addition, root hairs in PLDε-OE plants were more than twice the length of those of WT plants (Figure 7d,e), whereas root hair density did not differ between the genotypes. The effect on root hair length was apparent only under severe N-limiting conditions (0.1 mm N), and no difference in root hair length was observed at 6 or 60 mm N.

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Figure 6.  PLDε enhances biomass production at various levels of N. (a) Biomass accumulation. Four-week-old seedlings grown on MS salt agar plates were collected. Inset: biomass at 0.6 mm N. Values are means ± SD (= 30). (b) Root growth in response to N availability. Root length and number were measured 6 days after transferring 4-day-old seedlings to media containing MS salts and various N levels. To observe the effect of butanol on root growth and biomass accumulation, plants were grown on plates with or without 0.15% 1-butanol or 2-butanol for 4 weeks. MS salt medium with decreased P level was used to test the effect of P deprivation on growth. Values are means ± SD (= 30).

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Figure 7.  Primary root growth and root hair elongation under N-deprivation conditions. (a) Seedling morphology when seeds were germinated in MS salt containing 0.6 or 0.1 mm N. (b,c) Primary root length (10-day-old seedlings) and lateral root numbers. Values are means ± SD (= 20) for each of two independent experiments. (d) Root hair growth of seedlings germinated in growth medium containing 0.1 mm N for 7 days. Values are means ± SD (= 60). (e) Root hair length. Values are means ± SD (= 20). Asterisks indicate a significant difference at < 0.05, compared to WT, based on Student’s t test.

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PLDε-derived PA is involved in growth promotion

To determine whether PLD-produced PA is involved in growth alteration, we transferred Arabidopsis seedlings to growth media containing 1-butanol or 2-butanol. PLD uses 1-butanol, but not 2-butanol, as a substrate to form phosphatidylalcohol at the expense of PA. Thus, 1-butanol treatment was expected to suppress PLD-mediated PA production without inhibiting PLD degradation of membrane lipids. 1-butanol inhibited the number and length of lateral roots in all genotypes, but the magnitude of inhibition by 1-butanol was greater on OE plants and smaller on KO plants (Figure 6b). No significant difference was observed in the number and length of lateral roots among OE, WT and KO plants after 1-butanol treatment (Figure 6b). Treatment with 1-butanol also eliminated the difference in biomass accumulation among WT, OE and KO plants grown under high salinity (Figure 3h) and 6 mm N (Figure 6b). In contrast, the control treatment with 2-butanol had no inhibitory effect at the concentration tested (Figures 3h and 6b).

The level of PA in plants was measured to determine whether PA production was altered by KO and OE of PLDε. Leaf PA content from soil-grown KO plants was approximately 50% lower, and that in OE plants was 15% higher than in WT plants (Figure 8a). To measure PA changes in roots, seedlings were grown on plates with defined nitrogen levels. The level of PA in KO roots was only 67% of that in WT, whereas that in OE roots was slightly higher than that in WT at 2 mm N (Figure 8b). The levels of major membrane lipids, including PC, PG, monogalactosyl-diacylglycerol (MGDG) and digalactosyl-diacylglycerol (DGDG), were similar in KO, OE and WT roots. However, the PE level was higher in KO than in WT roots, but lower in OE than in WT roots. The inverse changes in PA and PE in roots suggest that most PA is derived from PLDε hydrolysis of PE (Figure 8c). The results from alcohol treatments and lipid analysis indicate that PLDε is active in PA production and that PLDε-produced PA is involved in growth promotion.

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Figure 8.  Effect of PLDε alterations on lipid content and composition. (a) Leaf PA content of 4-week-old, soil-grown plants under well-fertilized soil conditions. Values are means ± SE (= 5), and each replicate contained six leaves from six individual plants. The identification of the bars is the same as indicated in (b) and (c). (b) Lipid content in roots from seedlings grown in MS containing 2 mm N for 3 weeks. (c) Lipid molecular species of PE and PA in roots from seedlings grown in MS with 2 mm N for 3 weeks. Values are means ± SE (= 4), and each replicate contained at least 20 seedlings.

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Effect of PLDε alterations on nitrogen acquisition and assimilation

To determine the effect of PLDε alterations on N metabolism, we measured N uptake and metabolism under various levels of N. The expression of PLDε itself was induced twofold when seedlings were transferred from N-rich (60 mm) to N-limited (0.6 mm) conditions (Figure 9a, upper panel). Data from Genevestigator also indicated that expression of PLDε was increased under N-limited conditions, but deficiency in sulfur (S), potassium (K) or phosphorus (P) did not affect its expression (Figure 9b and Li et al., 2006). The expression of genes encoding two nitrate transporters, NRT1.1 and NRT2.1, which coordinate N absorption at different levels of N, were measured by real-time PCR. NRT1.1 is regarded as a dual-affinity nitrate transporter (Tsay et al., 1993; Huang et al., 1996), whereas NRT2.1 is a high-affinity, low-capacity nitrate transporter (Cerezo et al., 2001). The NRT1.1 mRNA level was high, whereas that of NRT2.1 was undetectable in all genotypes grown under high-N conditions. When seedlings were transferred from 60 to 0.6 mm N, expression of the high-affinity NRT1.1 in OE plants was higher than that in WT and KO plants (Figure 9a, lower panel). When seedlings were germinated and grown with limited N, the level of NRT1.1 expression was lowest in KO seedlings, and the level of NRT2.1 expression was highest in OE seedlings (Figure 9a). When nitrate uptake was compared in the genotypes, OE seedlings exhibited 1.5-fold higher nitrate uptake than WT and KO seedlings at 6 h after transferring the seedlings from a nitrate-starved to a 2 mm nitrate medium (Figure 9c). The changes in nitrate transporter expression and uptake suggest that OE of PLDε promotes nitrogen acquisition.

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Figure 9.  Changes in nitrate uptake and assimilation in PLDε-altered plants. (a) The expression levels of PLDε and nitrate transporters NRT1.1 and NRT2.1 under various N conditions was determined by real-time PCR using gene-specific primers (Table 1). Seedlings were grown in 60 or 0.6 mm N for 10 days, or 5-day-old seedlings grown on 60 mm N were transferred to 0.6 mm N for 5 days (60[RIGHTWARDS ARROW]0.6 mm). Values are means ± SD (= 3). (b) Expression of PLDε under normal and deficient nitrate (N), sulfur (S) and potassium (K) conditions. The data were obtained from Genevestigator (http://www.genevestigator.cthz.ch). (c) Nitrate uptake. Values are means ± SD (= 6). (d) Activity of enzymes involved in N assimilation and metabolism: NR (pmol NO2 per min per mg protein), NiR (pmol NH4+ per min per mg protein), GS (μmol γ-glutamylhydroxamate per min per mg protein) and GDH (μmol NADPH per min per mg protein). Ten-day-old seedlings grown on media containing nitrate at the concentraions shown were extracted for the assays. Values are means ± SD (= 3) from one of three independent experiments.

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The conversion of nitrate to amino acids requires several enzymes. The reductions of NO3 to NO2 and then to NH4+ are catalyzed sequentially by nitrate reductase (NR) and nitrite reductase (NiR). NH4+ is incorporated into organic molecules by the glutamine synthetase (GS)/glutamine synthase pathway (Crawford, 1995; Walch-Liu et al., 2006). The NR activity was similar in all the genotypes at 0.6 and 6 mm N, but OE seedlings showed a higher NiR activity than WT and KO seedlings at 0.6 and 6 mm N (Figure 9d). Similarly, the activity of GS was higher in OE seedlings (Figure 9d). In contrast, the activity of glutamate dehydrogenase (GDH), which catalyzes oxidative deamination of glutamate, was lower in OE than in WT and KO seedlings (Figure 9d). The difference in NiR, GS and GDH was greatest between OE and WT seedlings grown in low N (0.6 mm). The results indicate that OE of PLDε improves plant N utilization by enhancing the N assimilation pathway and decreasing glutamate catabolism.

Discussion

  1. Top of page
  2. Summary
  3. Introduction
  4. Results
  5. Discussion
  6. Experimental procedures
  7. Acknowledgements
  8. References

PLD and PA have been implicated in growth promotion through their roles in stimulating cellular growth regulators, such as mammalian target of rapamycin (mTOR), phosphoinositide-dependent kinase (PDK) and mitogen-activated kinases (Foster and Xu, 2003; Wang et al., 2006; Huang and Frohman, 2007), but direct evidence for promotion of growth at the organism level was lacking. The present study shows that PLDε promotes root growth and biomass accumulation. The increase results from both a bigger cell size and greater cell numbers. Arabidopsis has 12 PLD genes, but increased expression of other PLD genes, e.g. PLDα2, PLDα3 or PLDδ, does not result in overt growth enhancement (results not shown). The analysis of PLDε biochemical properties and expression provide insights into the distinctive effect of PLDε on plant growth. PLDε is the most permissive of all the characterized PLDs in terms of reaction requirements. However, the level of expression of PLDε in vegetative tissue is much lower than that of PLDα1. PLDε lacks a Ca2+–C2 interaction, making it less dependent on Ca2+ for its activation than other PLDs. In addition, the exclusive localization in membranes allows rapid activation of PLDε and access to membrane substrates without the relocation to the membrane that some of the other PLDs undergo. This could allow over-expressed PLDε to be active, resulting in production of PA that mediates the plant response to stimuli.

The results indicate that the growth promotion of PLDε is mediated via generation of the lipid messenger PA. The activity of PLDε in PA production has been demonstrated by biochemical assays of PLDε activity and by measuring altered PA levels in KO and OE plants. Lipid profiling of root tissue showed that most PA is derived from 34:3-PE and 34:2-PE, suggesting that unsaturated PA molecular species may be involved in sensing nutrient and osmotic cues to regulate root growth. In addition, the PA effect on growth alterations was corroborated by alcohol suppression of PA production. The increase in PA level is often transient (Wang et al., 2006), as PA is not metabolically stable. In addition, the location and timing of PA changes are important to PA functions. This may partly explain the results of direct measurements of PA in plants, which show that while KO of PLDε significantly decreases PA content, OE has a relatively small impact on the PA level in roots.

The potential for growth is affected greatly by an organism’s ability to sense and utilize available nutrients. Previously, PLDζs have been shown to play a role in the plant response to P deficiency by promoting primary root growth and membrane lipid remodeling as KO of PLDζ genes impedes root growth and P deficiency-induced lipid changes (Cruz-Ramirez et al., 2006; Li et al., 2006). The present data indicate that KO of PLDε had no overt effect on root growth and biomass accumulation under P-, K- or S-deprived conditions. However, PLDε-OE roots elongated faster, whereas KO roots elongated more slowly than WT roots. Opposite effects of PLDε KO versus OE on root elongation occurred in plants grown on agarose or agar media containing NO3:NH4+ (2:1) or NO3 only. The elongation of lateral roots in response to external N is regarded as an indicator of N signaling (Crawford, 1995; Walch-Liu et al., 2006; Hirel et al., 2007). The relationship between N availability, uptake and root development is well established, and root architecture and growth are critical to nutrient acquisition. The effects of PLDε on root growth and morphology are different at different levels of N. Under severe N deprivation (0.1 or 0.6 mm), PLDε promotes elongation of primary roots and root hairs, but no such effect was observed with sufficient N supply (6 or 60 mm). PLDε-OE plants exhibit increased expression of N transporters. Furthermore, greater differences in the levels of the nitrate assimilation enzymes NiR and GS were found between OE, KO and WT at 0.6 mm N than at 6 mm N. The results indicate that, under severe N deprivation, PLDε plays a role in increasing the root surface area to improve N uptake and utilization. These results are consistent with the notion that N uptake is one of the most critical N-utilization activities under N-limiting conditions in a number of plant species (Crawford, 1995; Hirel et al., 2007). At sufficient N supply, PLDε promotes lateral root growth and biomass production. The protein level and N content per unit biomass were not significantly different in WT and PLDε-altered plants. These results suggest that PLDε is involved in N signaling.

N is a vital nutrient for cellular structure and metabolism, and N deficiency is a major limiting factor that has an adverse impact on agricultural productivity. Considerable progress has been made in understanding the process of plant N uptake and assimilation in recent years (Crawford, 1995; Walch-Liu et al., 2006; Hirel et al., 2007). However, much less is known about signaling events in plant response to N availability. The present finding of the growth effect of PLDε raises intriguing questions as to whether PLD and PA-mediated signaling play a role in connecting the sensing of external nutrient cues at membranes to translational regulation and growth alterations. PA has been found to bind to PDK1. The PA–PDK1 interaction activates AGC2-1 kinase to promote root hair growth (Anthony et al., 2004). This is consistent with the present results showing that over-expressing PLDε promotes root hair growth. PDK1 also phosphorylates S6K (Mahfouz et al., 2006). In mammalian cells, S6K is a nodal point integrating nutrient and stress inputs through translational activity to regulate growth and nutrient usage (Wullschleger et al., 2006). Over-expression of PLD1 increases S6K activity and cell size, whereas suppression of PLD1 reduces cell size (Fang et al., 2003). PA has been shown to bind to S6K and its upstream kinase mTOR in mammalian cells. The PA–mTOR interaction occurs at a site that competes with rapamycin binding (Fang et al., 2001; Lehman et al., 2007). The effect of PA on AtTOR is unknown, but plant TOR is insensitive to rapamycin and thus may not bind rapamycin (Mahfouz et al., 2006). It would be of great interest to determine whether PA interactions with multiple kinases may tether these proteins to membranes, and produce a multi-protein signaling complex involved in growth regulation.

The enhancement of growth by PLDε also occurs under hyper-osmotic stress imposed by high salinity and water deficiency (Figure 3). Activation of PLD has been found to occur under various hyper-osmotic conditions (Wang, 2004; Testerink and Munnik, 2005). In nature, simple hyper-osmotic stress does not occur because of the interplay between water availability and nutrient transport. It is known that water deficiency impedes nutrient uptake and that drought induces N deficiency (Heckathorn et al., 1997; Foyer et al., 1998). Thus, the altered growth in PLDε-OE and KO plants under hyper-osmotic stress could be related to the positive role of PLDε and PA in nutrient signaling. These results raise the exciting possibility that PLD and PA-based membrane lipid signaling acts as a key integrator of multiple plant stresses for optimal cell growth in response to drought and nutrient stresses, and this calls for further investigation.

Experimental procedures

  1. Top of page
  2. Summary
  3. Introduction
  4. Results
  5. Discussion
  6. Experimental procedures
  7. Acknowledgements
  8. References

Isolation of PLDε knockout and HA-tagged PLDs expressed in plants

A PLDε T-DNA insertion mutant was identified from SALK_023603 of the Salk Arabidopsis T-DNA knockout collection from the Ohio State University Arabidopsis Biological Resource Center. The homozygous T-DNA insertion mutant pldε-1 was isolated by PCR-based screening using PLDε-specific primers and a T-DNA left border primer (Table 1). The loss of transcription of PLDε was confirmed by RT-PCR using PLDε-specific primers. To over-express PLDε in Arabidopsis plants, a DNA fragment for the PLDε gene was amplified from wild-type Columbia ecotype Arabidopsis genomic DNA using gene-specific primers (Table 1). PLDε was fused with an HA tag at the 3′ end, and then cloned into a binary vector pKYLX7 under the control of the CaMV 35S promoter.

Table 1.   Primers used for mutant isolation, cloning and real-time PCR
PurposeNamePrimer sequences
PLDε cloningPLDεForward 5′-AGA GGG ATC CAT GGA GCT TGA AGA ACA GAA GAA G-3′ Reverse 5′-GTT AGG CCT GGT GGT TAG AAC AGG AGG AAA CA-3′
Knockout screeningPLDεForward 5′-AGA GGG ATC CAT GGA GCT TGA AGA ACA GAA GAA G-3′ Reverse 5′-GTT AGG CCT GGT GGT TAG AAC AGG AGG AAA CA-3′
Left border5′-GCG TGG ACC GCT TGC TGC AAC T-3′
Reverse transcription and real-time PCRPLDεForward 5′-TAT CTT GAA CCG GGA TGG TGC AGA-3′ Reverse 5′-TAG GGT TTA GTG CCC ATC CTG CAA-3′
ComplementationPLDεForward 5′-GAA TTC GCG TCC TCG CAT GTC TCA GGT AAA-3′ Reverse 5′-GGA TCC TGC CCT CAT GTG TTC TTA TCA AGG ACA-3′
Complementation confirmationTeasy AscForward 5′-ATG GCG CGC CAT ATG GTC GAC CTG CAG-3′ Reverse 5′-ATG GCG CGC CCG ACG TCG CAT GCT C-3′
Real-time PCRNRT1.1Forward 5′-ACG TCT TAG AAC TGC ACA CGC TCA-3′ Reverse 5′-AGA TTA ACG CTT CGC CGA TAC CGA-3′
NRT2.1Forward 5′-CTT TGT ACC CGG TTG GTT GCA CAT-3′ Reverse 5′-TTT GTC TTT GGC AAC TTC TCC CGC-3′
UBQ10Forward 5′-CAC ACT CCA CTT GGT CTT GCG T-3′ Reverse 5′-TGG TCT TTC CGG TGA GAG TCT TCA-3′

Plant growth and treatments

Surface-sterilized seeds were germinated in MS salt agar or agarose (to remove potential nutrient contaminants) for 4 days, and then transferred to modified MS salt agar plates containing 0.1, 0.6, 2, 6 or 60 mm N (NO3:NH4+; 2:1) or NO3 only. KCl was added to compensate for the lower than normal K+ concentration in media with reduced KNO3 levels (Martin et al., 2002). Seedlings were grown on plates in a vertical orientation in a growth chamber with a 16 h light/8 h dark cycle, 23/21°C, under cool fluorescent white light (200 μmol m−2 sec−1). Alternatively, seeds were directly germinated on agar plates and grown without a transfer step. For osmotic stress experiments, surface-sterilized seeds were germinated on or 4-day-old seedlings were transferred to MS salt growth media containing 50 or 100 mm NaCl, 5 or 8% PEG, or 100 mm sorbitol. For experiments on soil-grown plants, plants were grown in growth chambers with a 12 h light/12 h dark cycle, 23/21°C, 50% humidity, at 200 μmol m−2 sec−1 of light intensity, and watered with fertilizer (15-5-15 Cal-Mag, The Scotts Company, http://www.scottspro.com, 200 ppm nitrogen) once a week (well-fertilized conditions) or only once per life cycle (less well-fertilized). The fertilizer contained 15% total nitrogen (1.2% ammonia, 11.75% nitrate, 2.05% urea), 5% available phosphate and 15% soluble potassium.

RNA extraction and real-time PCR

Total RNA was extracted from leaves or seedlings using a CTAB method. DNA was removed from RNA by digestion with RNase-free DNase I (Li et al., 2006). RNA without DNA contamination was used as a template for RT-PCR for synthesis of cDNA using an iScript kit (Bio-Rad, http://www.bio-rad.com/). Quantitative real-time PCR was performed using a MyiQ sequence detection system (Bio-Rad) by monitoring SYBR green fluorescent labeling of double-stranded DNA synthesis as described previously (Li et al., 2006). The efficiency of cDNA synthesis was assessed by real-time PCR amplification of a control gene encoding UBQ10 (At4g05320; primer sequences in Table 1); the Ct value for the UBQ10 gene was 20 ± 0.5. Only cDNA preparations that yielded similar Ct values for the control gene were used for determination of PLD gene expression. The level of PLD expression was normalized to that of UBQ10 by subtracting the Ct value of UBQ10 from that of the PLD genes (Li et al., 2006).

Subcellular fractionation, PLDε–HA purification and PLD activity assays

Proteins were extracted from leaves of 4-week-old plants using chilled buffer A (Qin et al., 1997), followed by centrifugation at 6000 g for 10 min. The supernatant was centrifuged at 100 000 g for 60 min, and the resultant supernatant and pellet are referred to as the soluble and microsomal fractions, respectively. The microsomal fraction was separated into plasma membrane and intracellular membrane fractions by two-phase partitioning as described previously (Fan et al., 1999). Marker enzymes for the plasma membrane and intracellular membranes are ATPase and cytochrome c reductase, respectively (Fan et al., 1999). PLDε was purified from PLDε-OE Arabidopsis leaves using HA antibody affinity chromatography. Briefly, protein extracts were incubated with HA monoclonal antibody (1:500) at 4°C for 2 h. Protein A Agarose was added to the mixture and incubated with agitation for 2 h at 4°C. Beads were pelleted at 6000 g and washed four times for 3 min each with a wash buffer containing 0.5% Triton X-100. The purified PLDs were assayed under the activity conditions previously defined for PLDα1, β, δ and ζ1 (Qin et al., 1997; Pappan et al., 1998; Wang and Wang, 2001; Qin and Wang, 2002). To assay substrate usage, fluorescent 1-Oleoyl-2-[12[(7-nitro-2-1,3-benzoxadiazol-4-yl) amino]dodecanoyl]-sn-Glycero-3-PC, -PE, -PG, -PS were used, and each reaction contained PLDε–HA isolated from 1 mg leaf proteins under the PLDα1 reaction conditions (50 mm Ca2+, 0.5 mm SDS, 2 mm lipids). The resultant lipids were separated on TLC plates and quantified by fluorescence spectrophotometry (excitation at 460 nm, emission at 534 nm) (Pappan et al., 1998).

Subcellular localization of PLDε–YFP

Arabidopsis PLDε cDNA was cloned by PCR amplification from an Arabidopsis leaf cDNA RACE pool. The amplified PCR product was directly cloned into p35S_FAST/YFP, which was derived from p35S_FAST (Ge et al., 2005) by introducing eYFP (Sigma, http://www.sigmaaldrich.com/). Agro-infiltration for transient protein expression in tobacco leaves was performed as described by Voinnet et al. (2003). In brief, Agrobacterium tumefaciens strain C58C1 carrying binary constructs was grown to stationary phase at 28°C in YEP medium. Bacterial cells were collected by centrifugation at 6000 g for 15 min at room temperature, and resuspended in 10 mm MES (pH 5.7), 10 mm MgCl2 and 150 mg ml−1 acetosyringone. For co-infiltration, Agrobacterium cultures carrying various constructs were mixed at an equal ratio and left for 3 h at room temperature before infiltration. Leaves of 3-week-old Nicotiana benthamiana plants were infiltrated with the bacterial cultures through abaxial air spaces. In all experiments, Agrobacterium C58C1 carrying the 35S:p19 construct (Voinnet et al., 2003) was co-infiltrated to achieve a maximum level of protein expression. The YFP fluorescence was observed under a Zeiss LSM 510 confocal/multi-photon microscope (http://www.zeiss.com/).

Immunoblotting of PLD

Proteins were extracted from leaves or seedlings as previously described (Qin et al., 1997). Homogenates were centrifuged at 6000 g for 10 min. For PLD–HA detection, supernatant proteins (30 μg per lane) were separated by 8% w/v SDS–PAGE, followed by transfer to a PVDF membrane. Membranes were blotted with anti-HA antibody (1:1000) and then incubated with a secondary antibody as described previously (Zhang et al., 2004).

Leaf cell size and lipid analysis

Leaf discs (0.5 cm diameter) were taken from the middle of fully expanded leaves from 5-week-old plants, and fixed in ethanol:glacial acetic acid (3:1 v/v) for 30 min. Leaf discs were transferred sequentially to 75, 50, 25 and 0% ethanol (v/v) for 15 min each. Cell sizes were measured under a microscope using IMAGEPRO software (Media Cybernetics, http://www.mediacy.com). Lipids were extracted and analyzed by ESI-MS/MS, and the levels of PA, PC, PE, PG, phosphatidylinositol (PI) and PS molecular species were added to determine lipid content for each head-group class as previously described (Devaiah et al., 2006).

Nitrate uptake and determination of levels of assimilation enzymes

Nitrate uptake was assayed as described by Doddema and Telkamp (1979). To measure the levels of N-metabolizing enzymes, 10-day-old seedlings were homogenized with chilled extraction buffer (100 mm K2HPO4 pH 7.5, 1 mm DTT, 1 mm EDTA and 10 mm cysteine), followed by centrifugation at 10 000 g at 4°C for 15 min. The supernatant was used to assay the activities of NR (Wilkinson and Crawford, 1991), NiR (Takahashi et al., 2001), GS (Rhodes et al., 1975) and GDH (Turano et al., 1996), as previously described.

Acknowledgements

  1. Top of page
  2. Summary
  3. Introduction
  4. Results
  5. Discussion
  6. Experimental procedures
  7. Acknowledgements
  8. References

This work was supported by grants from the National Science Foundation (IOS 0818740) and the US Department of Agriculture (2007-35318-18393). Lipid analysis at Kansas Lipidomics Research Center was supported by the National Science Foundation (MCB 0455318, DBI 0521587), the Kansas Technology Enterprise Corporation, and the K-IDeA Networks of Biomedical Research Excellence (INBRE) of the National Institutes of Health (P20RR16475). We thank Mary Roth for technical assistance. We thank David Baulcombe for providing vectors of p19 and pCH32, and Yiji Xia for the binary vector p35S_FAST.

References

  1. Top of page
  2. Summary
  3. Introduction
  4. Results
  5. Discussion
  6. Experimental procedures
  7. Acknowledgements
  8. References