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PHOSPHATIDIC ACID PHOSPHOHYDROLASE1 and 2 Regulate Phospholipid Synthesis at the Endoplasmic Reticulum in Arabidopsis

Peter J. Eastmond, Anne-Laure Quettier, Johan T.M. Kroon, Christian Craddock, Nicolette Adams, Antoni R. Slabas

Published August 2010. DOI: https://doi.org/10.1105/tpc.109.071423

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Abstract

Phospholipid biosynthesis is essential for the construction of most eukaryotic cell membranes, but how this process is regulated in plants remains poorly understood. Here, we show that in Arabidopsis thaliana, two Mg2+-dependent phosphatidic acid phosphohydrolases called PAH1 and PAH2 act redundantly to repress phospholipid biosynthesis at the endoplasmic reticulum (ER). Leaves from pah1 pah2 double mutants contain ~1.8-fold more phospholipid than the wild type and exhibit gross changes in ER morphology, which are consistent with massive membrane overexpansion. The net rate of incorporation of [methyl-14C]choline into phosphatidylcholine (PC) is ~1.8-fold greater in the double mutant, and the transcript abundance of several key genes that encode enzymes involved in phospholipid synthesis is increased. In particular, we show that PHOSPHORYLETHANOLAMINE N-METHYLTRANSFERASE1 (PEAMT1) is upregulated at the level of transcription in pah1 pah2 leaves. PEAMT catalyzes the first committed step of choline synthesis in Arabidopsis and defines a variant pathway for PC synthesis not found in yeasts or mammals. Our data suggest that PAH1/2 play a regulatory role in phospholipid synthesis that is analogous to that described in Saccharomyces cerevisiae. However, the target enzymes differ, and key components of the signal transduction pathway do not appear to be conserved.

INTRODUCTION

Phospholipids are the major component of most eukaryotic cell membranes, and these membranes play a fundamental role in compartmentalizing the biochemistry of life. The quantity and composition of phospholipids are tightly regulated so that membranes can maintain their structure and function, despite developmental and environmental changes (Ohlrogge and Browse, 1995). Research performed in mammals and yeast (Saccharomyces cerevisiae) has shown that metabolite sensing and signaling mechanisms are critical for this regulation. These mechanisms function at the transcriptional or posttranslational level, allowing the cell to sense changes in the availability of key lipid intermediates and adjust phospholipid synthesis (and turnover) accordingly (Rawson, 2003; Jackowski and Fagone, 2005; Kent, 2005; Carman and Henry, 2007; Sugimoto et al., 2008; Carman and Han, 2009). It has been suggested that analogous signaling mechanisms are also likely to exist in plants (Xu et al., 2005), but they have yet to be elucidated.

In yeast, a transcriptional mechanism has recently been described that coordinately regulates phospholipid biosynthetic gene expression in response to changes in the level of phosphatidic acid (PA), which is a known signaling compound and precursor for all phospholipids (Carman and Henry, 2007). Many yeast genes that encode phospholipid biosynthetic enzymes contain upstream activating sequence inositol-responsive (UASINO) elements in their promoters. Their expression is enhanced by interaction of the Ino2p-Ino4p transcription factor complex with UASINO elements, whereas it is blocked by interaction of the repressor protein OVERPRODUCER OF INOSITOL1 (Opi1p) with Ino2p (Carman and Henry, 2007). PA has a crucial role in the Opi1p-mediated repression of gene expression because it tethers Opi1p at the nuclear–endoplasmic reticulum (ER) membrane, together with a vesicle-associated protein homolog called SUPRESSOR OF CHOLINE SENSITIVITY2 (Loewen et al., 2004). A reduction in PA concentration promotes translocation of Opi1p from the nuclear-ER membrane into the nucleus, where it interacts with Ino2p to repress gene expression and reduce the rate of phospholipid synthesis (Loewen et al., 2004).

In yeast, cellular quantities of PA can be regulated by the activity of the Mg2+-dependent PA phosphohydrolase Pah1p (Han et al., 2006). Mutants in the PAH1 gene have elevated levels of PA and show massive expansion of the nuclear-ER membrane and abnormally high expression of key phospholipid biosynthetic genes (Santos-Rosa et al., 2005; Han et al., 2006, 2007). Conversely, overexpression of PAH1 leads to repression of phospholipid biosynthetic gene expression, and this response depends upon Pah1p catalytic function (O'Hara et al., 2006; Han et al., 2007). Pah1p activity is regulated via phosphorylation by cyclin-dependent kinase Cdc28p (also known as Cdk1p) and dephosphorylation by a Nem1p-Spo7p phosphatase complex situated on the nuclear-ER membrane (Santos-Rosa et al., 2005; O'Hara et al., 2006). In yeast, the phosphorylation status of Pah1p can therefore coordinate phospholipid biosynthetic gene expression and growth of the nuclear-ER membrane with cell cycle progression (O'Hara et al., 2006; Han et al., 2007).

In addition to playing a regulatory role through the modulation of PA levels, yeast Pah1p also provides substrate for the production of membrane and storage lipids (Carman and Henry, 2007). The yeast pah1Δ mutant is severely compromised in triacylglycerol (TG) production, particularly during stationary phase when TG normally accumulates (Han et al., 2006). Disruption of a mammalian PAH1 homolog called Lipin-1 also causes a reduction in adipose tissue mass (Péterfy et al., 2001), and Lipin-1 can complement the yeast pah1Δ mutant (Han et al., 2006; Grimsey et al., 2008). Yeast Pah1p and mammalian Lipin-1 reside in the cytosol and are thought to translocate to the ER membrane to bind PA (Han et al., 2006; Péterfy et al., 2010). However, Lipin-1 has also been found in the nucleus (Péterfy et al., 2001). Finck et al. (2006) reported that Lipin-1 is a bifunctional protein that has a transcriptional role in mammals that is independent of its catalytic activity. Lipin-1 functions as a transcriptional coactivator by interacting directly with the nuclear hormone receptor PPARα, leading to the formation of a complex that can modulate the expression of multiple genes involved in lipid metabolism (Finck et al., 2006; Donkor et al., 2009).

Plants contain multiple PA phosphohydrolase isoforms, and the metabolic function of these enzymes has been the focus of a number of studies (Joyard and Douce, 1979; Block et al., 1983; Pearce and Slabas, 1998; Pierrugues et al., 2001; Nakamura et al., 2007; Nakamura et al., 2009). However, it has yet to be established whether any of these proteins play a regulatory role in plant lipid metabolism, either directly or indirectly. Arabidopsis thaliana contains two genes (At PAH1 and At PAH2) that are similar to yeast PAH1 (Han et al., 2006) and mammalian Lipin-1 (Péterfy et al., 2001). Nakamura et al. (2009) recently studied these genes and proposed that they play a metabolic role in the provision of substrate for galactolipid biosynthesis via the eukaryotic pathway. Two biosynthetic pathways exist in plants, namely, the prokaryotic pathway situated entirely in the chloroplast and the eukaryotic pathway that proceeds via the ER (Ohlrogge and Browse, 1995). In Arabidopsis, both pathways contribute approximately equally to the synthesis of galactolipids (Ohlrogge and Browse, 1995). Here, we show that a pah1 pah2 double mutant not only exhibits a reduction in galactolipids derived from the eukaryotic pathway but also a dramatic and disproportionate increase in the level of phospholipids, coupled with overexpansion of the ER membrane. Phosphatidylcholine (PC) is the major phospholipid in plants (Ohlrogge and Browse, 1995), and the net rate of incorporation of [methyl-14C]-choline into PC is increased ~1.8-fold in the pah1 pah2 double mutant. Analysis of transcript abundance suggests that genes encoding several key enzymes of phospholipid synthesis are upregulated. Our data suggest that PAH1 and PAH2 function indirectly as repressors of phospholipid biosynthesis in Arabidopsis and this regulation occurs at least partly at the level of transcript abundance for multiple enzymes.

RESULTS

Arabidopsis Contains Two Mg2+-Dependent Phosphatidate Phosphohydrolases

Arabidopsis contains two genes, which are similar to yeast PAH1 (Han et al., 2006) and human Lipin-1 (Péterfy et al., 2001). Nakamura et al. (2009) recently designated these loci At PAH1 and At PAH2. The open reading frames for both genes were amplified by PCR from Arabidopsis cDNA. The PAH1 nucleotide sequence is 2742 bp long and encodes a 904–amino acid polypeptide. PAH2 is 2793 bp long and encodes a 930–amino acid polypeptide. PAH1 and PAH2 exhibit very little sequence similarity to yeast Pah1p and human Lipin-1 with the exception of conserved N- and C-terminal regions (Péterfy et al., 2001; Nakamura et al., 2009; Figure 1A). The C-terminal region contains a haloacid dehalogenase superfamily (HAD-like) domain with the catalytic motif DI/VDGT, which is characteristic of all the Mg2+-dependent PA phosphatases that have been identified to date (Han et al., 2006; Figure 1B). The precise function of the conserved N-terminal region is not known.

Figure 1.
Figure 1.

Arabidopsis Contains Two Homologs of Yeast PAH1 That Are Expressed in Many Tissues and Are Located in the Cytosol.

(A) A schematic diagram showing the positions of the conserved N-terminal (red) and C-terminal (blue) regions in Arabidopsis PAH1 and PAH2, yeast Pah1p, and human Lipin1. aa, amino acids.

(B) An amino acid sequence alignment surrounding the HAD-like domain in the C-terminal regions of Arabidopsis PAH1 and PAH2, yeast Pah1p, and human Lipin-1. The alignment was performed using ClustalX (version 1.83). The conserved catalytic motif DI/VDGT is boxed.

(C) Real-time PCR analysis of the relative expression of PAH1 and PAH2 in various tissues of Arabidopsis. Torpedo (TO), walking stick (WS), curled cotyledon (CC), green cotyledon (GC), and mature seed (MS) describe seeds at different stages in development. SE, LE, RO, ST, and FL are seedling, leaf, root, stem, and flower, respectively. Values are the mean ±se of measurements on RNA extractions from four separate batches of tissue. Equal quantities of total RNA were used for each sample.

(D) Confocal images of agrobacterium-infiltrated N. benthamiana leaf cells expressing GFP-PAH1 and GFP-PAH2. Bars = 10 μm.

Analysis of relative PAH1 and PAH2 transcript abundance by real-time PCR showed that both genes are expressed in all the tissues, organs, and stages in development that were investigated but that transcript abundance for both genes is highest in seeds during the maturation stage (Figure 1C). These findings are consistent with public Arabidopsis microarray data for At3g09560 and At5g42870 available at https://www.genevestigator.ethz.ch (Zimmermann et al., 2004). Subcellular localization prediction software suggests that neither protein has transit peptides or transmembrane regions, and subcellular fractionation combined with immunoblotting has indicated that the proteins are most likely to be cytosolic (Nakamura et al., 2009). To investigate where PAH1 and PAH2 are located in vivo, N-terminal green fluorescent protein (GFP) fusions were transiently expressed in Nicotiana benthamiana leaves under the control of the 35S promoter. Both fusion proteins were found to be located predominantly in the cytosol (Figure 1D). No signal was detected in the nucleus or chloroplast.

To investigate the enzymological properties of PAH1 and PAH2, His6-tagged proteins were heterologously expressed in Escherichia coli and purified, and their identities were confirmed by matrix-assisted laser desorption/ionization-time of flight mass spectrometry (see Supplemental Figure 1 online). The activities of purified recombinant PAH1 (rPAH1) and PAH2 (rPAH2) were determined using a radiolabel PA phosphatase assay method, adapted from Han et al. (2006), in which the substrate forms a mixed micelle with Triton X-100 in a molar ratio of 10:1 (Carman et al., 1995). The concentration of PA used in this study was 3 mM. Values higher than 5 mM have been shown to inhibit the activity of yeast Pah1p (Lin and Carman, 1989). Both rPAH1 and rPAH2 were able to dephosphorylate 32P-labeled PA in a manner that was proportional with respect to time (Figure 2A) and to the amount of enzyme added to the assay (Figure 2B). The specific activity of rPAH2 appears to be greater than that of rPAH1 (Figure 2A). However, it is not known whether this would be true of the native proteins expressed in Arabidopsis. The pH optimum for both enzymes was 6.5 (Figure 2C). In comparison, the pH optimum for recombinant yeast Pah1p is 7.5 (Han et al., 2006). PA phosphatase activity exhibited a dose-dependent requirement for MgCl2, with the maximum activity of rPAH1 and rPAH2 attained at a final concentration of 2 and 1 mM, respectively (Figure 2D). No activity was observed in the presence of MgCl2 plus 5 mM EDTA or when MgCl2 was substituted with 2 mM MnCl2 or 2mM HgCl2. The activities were not sensitive to the thioreactive compound N-ethylmaleimide (2 mM), and 32P-labeled lyso-PA was not dephosphorylated by either protein (data not shown). These properties are generally consistent with those reported for yeast Pah1p (Han et al., 2006).

Figure 2.
Figure 2.

Expression of PAH1 and PAH2 in E. coli and Characterization of PA Phosphohydrolase Activity.

The effects of time (A), enzyme concentration (B), pH (C), and Mg2+ concentration (D) on 32P-PA hydrolysis by recombinant Arabidopsis PAH1 (closed circles) and PAH2 (open circles). Values are the mean of three separate assays and standard errors were <5% of the mean.

PAH 1 and 2 Function Redundantly and Are Required for Normal Vegetative Growth

To investigate the physiological function of PAH1 and PAH2, T-DNA insertion mutants were obtained from T-DNA express (Alonso et al., 2003) and from Institut National de la Recherche Agronomique (INRA) (Brunaud et al., 2002). Homozygous mutant lines were identified for each mutant allele, and the position of the insertion sites was confirmed by PCR amplification and sequencing of genomic DNA flanking both borders (Alonso et al., 2003). The pah1 (SALK_042970) mutant contains a T-DNA inserted in the first exon of At3g09560, at position +576 bp relative to the start codon (Figure 3A). The pah2-1 (SALK_047457), pah2-2 (FLAG_076D10), and pah2-3 (FLAG_196D12) mutants contain T-DNA inserted in the splice site between the seventh intron and eighth exon, in the fourth intron and in the second exon of At5g42870, at positions +3201, +2589, and +1105 bp, respectively (Figure 3A).

Figure 3.
Figure 3.

Isolation of pah1 and pah2 Mutants.

(A) A schematic diagram to show the positions of T-DNA insertions in pah1 and pah2 mutant alleles. Insertions are numbered relative to the initiation codon. Black bars are exons, and the white bars and arrows indicate the 5′ and 3′UTRs, respectively. The position of primers used in (B) is indicated by arrows.

(B) RT-PCR analysis of PAH1 and PAH2 transcripts in the pah1 and pah2-1 mutants. The position of primers is shown in (A). ACT2 was used as a positive control for the RT-PCR. Single PCR reactions were performed on RNA from individual plants of each genotype. WT, wild type.

(C) to (E) Images of rosettes (C), individual leaves (D), and siliques (E) of wild-type, single, and double pah1 pah2-1 mutant plants. Bars = 3, 1, and 1 cm, respectively. Plants were grown on soil in a growth chamber set to 21°C 16 h light/18°C 8 h dark; PPFD = 150 μmol m−2 s−1.

(F) Image of wild-type and pah1 pah2-1 seedlings grown for 5 d in the light on half-strength Murashige and Skoog (MS) salts plus 1% (w/v) sucrose. Bar = 1 cm.

Examination of all of the homozygous pah1 and pah2 alleles revealed no obvious phenotypic abnormalities. However, since PAH1 and PAH2 appear to be expressed in all plant tissues (Figure 1C), it remained conceivable that the two genes function redundantly. To investigate this possibility, double mutants were generated between pah1 and the pah2-1, pah2-2, and pah2-3 alleles. Phenotypic analysis of these three double mutants showed that they all have an abnormal growth phenotype that is recessive and cosegregates with the presence of T-DNA insertions in PAH1 and PAH2 (see Supplemental Table 1 online). Detailed studies were performed on pah1 pah2-1, which has a common genetic background (ecotype Columbia-0). This double mutant is the same combination of alleles as that described recently by Nakamura et al. (2009). PCR experiments performed on cDNA from homozygous pah1 pah2-1 plants showed that expression of wild-type PAH1 and PAH2 mRNA is disrupted in this line (Figure 3B).

In terms of gross morphology, pah1 pah2-1 plants grow more slowly than the wild type in controlled conditions (21°C and 16 h light, 18°C and 8 h dark; PPFD = 150 μmol m−2 s−1), producing stouter rosette leaves with shorter petioles (Figures 3C and 3D). They also have shorter siliques (Figure 3E), seedling primary roots (Figure 3F), and primary racemes as well as reduced seed set (Table 1). However, the average weight of individual mature seeds produced by pah1 pah2-1 plants was not significantly different from the wild type (Table 1) and their morphology appeared normal. Furthermore, the fact that pah1 pah2-1 plants produced shorter siliques (Table 1), containing less seed, was not the result of poor fertility or embryo abortion. Short siliques were still obtained when pah1 pah2-1 plants were fertilized with wild-type pollen (12.5 ± 0.2 mm, n = 8, ±se), and inspection of developing pah1 pah2-1 siliques on self-fertilized plants suggested that the frequency of embryo abortion was <1% and therefore no more common than in the wild type.

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Table 1.

Phenotypic Analysis of the pah1 pah2-1 Double Mutant

To investigate whether the reduced growth rate of pah1 pah2-1 plants could be caused by a defect in net CO2 fixation, a number of parameters associated with photosynthetic capacity were measured (Table 2). The apparent rate of CO2 assimilation, expressed on a per unit leaf area basis, was slightly greater in pah1 pah2-1 leaves relative to the wild type, under both ambient and saturating CO2 conditions (Table 2). The total amount of chlorophyll a and b, per unit fresh weight (FW), was also increased in pah1 pah2-1 leaves (Table 2). However, the maximum quantum efficiency (Fv/Fm) of photosystem II and the chlorophyll a/b ratio were not significantly changed (Table 2). These data suggest that the growth phenotype of pah1 pah2-1 plants is unlikely to be explained by a defect in the chloroplast photosynthetic apparatus.

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Table 2.

Analysis of Photosynthetic Parameters in pah1 pah2-1 Leaves

pah1 pah2 Double Mutants Overproduce Phospholipids

A preliminary analysis of whether lipid metabolism was altered in different tissues of pah1 pah2-1 plants was performed by measuring the fatty acid content and composition of leaves, roots, and seeds. Analysis of pah1 pah2-1 seeds suggested that the total fatty acid content was ~15% lower than the wild type, on a per seed basis, and that the reduction is due primarily to less octadecadienoic acid (18:2), octadecatrienoic acid (18:3), and eicosenoic acid (20:1) (Figure 4A). Approximately 95% of the fatty acids present in wild-type seeds are found in TG (Ohlrogge and Browse, 1995), and extraction and quantification of TG confirmed that this is also true of pah1 pah2-1 seeds (95.4% ± 3.1%, n = 10, ±se). When the total fatty acid content of pah1 pah2-1 roots and rosette leaves was measured, the double mutant was found to contain up to 80 and 20% more total fatty acids, respectively, on a per unit FW basis (Figures 4B and 4C). In leaves, the levels of hexadecanoic acid (16:0), octadecenoic acid (18:1), 18:2, and 18:3 were all increased, while the levels of hexadecenoic acid (16:1) and hexadecatrienoic acid (16:3) were not significantly changed (Figure 4C). In wild-type Arabidopsis leaves, 16:1 and 16:3 are almost exclusively found in phosphatidylglycerol and monogalactosyldiacylglycerol (MGD), respectively, which are chloroplast membrane lipids synthesized predominantly by the prokaryotic pathway (Ohlrogge and Browse, 1995). Assuming that the distribution of these fatty acids within the lipid classes is not greatly changed in pah1 pah2-1 leaves, the increase in total fatty acid levels seen in Figure 4C could be explained by net overproduction of ER-derived lipids, synthesized via the eukaryotic pathway.

Figure 4.
Figure 4.

The Fatty Acid Content and Composition of Tissues from pah1 pah2-1 Double Mutant Plants.

The fatty acid content and composition of seeds (A), roots (B), and leaves (C). Values are the mean ± se of measurements made on samples from 5 to 10 different plants of each genotype and are expressed on a per seed basis in (A) and on a per unit FW basis in (B) and (C). *Significantly different from the wild type (WT; P < 0.05).

Quantification of polar lipid classes was performed on leaf and root extracts using two-dimensional thin layer chromatography (TLC) separation followed by gas chromatography (GC) analysis of the methyl esters. This analysis showed that the amount of PC, phosphatidylethanolamine (PE), phosphatidylinositol, and PA, on a per mg FW basis, are all increased in pah1 pah2-1 double mutant leaves by between 70 and 100%, while MGD and digalactosyldiacylglycerol (DGD) exhibit a comparatively small decrease in absolute abundance of around 5 and 25%, respectively (Figure 5A). In roots, where galactolipids normally only make up a very minor proportion of the total polar lipids, a 70 to 100% increase in the amount of phospholipids was also observed in pah1 pah2-1 (Figure 5B). The increase in the absolute quantity of PC in leaves and roots was also observed using an enzyme-coupled fluorometric assay method described by Nanjee et al. (1991). Using this method, there was a 1.72- and 1.78-fold increase in the amount of PC in pah1 pah2-1 leaves and roots, respectively (n = 5, se < 0.3).

Figure 5.
Figure 5.

Polar Lipid Measurements on Tissues from pah1 pah2-1 Double Mutant Plants.

The absolute quantity of different polar lipid classes from leaves (A) and roots (B) determined following separation by TLC and quantification by GC of fatty acid methyl esters. Values are the mean ± se of measurements made on samples from four different plants for each genotype and are expressed on a per unit FW basis. *Significantly different from the wild type (WT; P < 0.05). PI, phosphatidylinositol; PS, phosphatidylserine; SL, sulfolipids; PG, phosphatidylglycerol.

The fatty acid composition of the lipid classes was altered relatively little in pah1 pah2-1 leaves (Table 3). However, a small increase in the mol % of 16:3 and decrease in 18:3 were observed in MGD (Table 3). A relative decrease in DGD content and a shift in the ratio of 16:3 to 18:3 in MGD have also been observed by Nakamura et al. (2009) in pah1 pah2-1 and are consistent with a reduction in the contribution of the eukaryotic pathway to galactolipid synthesis. The fatty acid composition of the PA in pah1 pah2-1 leaves was found to be quite similar to that of PC (Table 3). Furthermore, positional analysis of the fatty acids suggested an enrichment of 18 carbon fatty acids at the sn-2 position on the glycerol backbone (89.2 ± 7 mol %, n = 3, ±se). These data are consistent with the PA originating from the eukaryotic pathway (Heinz and Roughan, 1983; Xu et al., 2005). A number of Arabidopsis mutants that are impaired in trafficking of lipids to the chloroplast for galactolipid synthesis have been shown to accumulate TG and trigalactosyldiacylglycerol in their leaves (Xu et al., 2005; Awai et al., 2006). However, using standard TLC methods described by Xu et al. (2005), neither compound was detected in significant quantities in leaves from wild-type or pah1 pah2-1 plants.

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Table 3.

Fatty Acid Composition of Polar Lipid Classes from Leaves of the Wild Type and the pah1 pah2-1 Double Mutant

pah1 pah2 Double Mutants Exhibit Overexpansion of the ER Membrane

In yeast, disruption of PAH1 leads to massive expansion of the nuclear-ER membrane and greatly enlarged nuclei (Santos-Rosa et al., 2005; Han et al., 2007). To investigate whether this is also the case in Arabidopsis, nuclear-, ER lumen–, and ER membrane–targeted GFP (Irons et al., 2003) were transiently expressed in pah1 pah2-1 and wild-type Arabidopsis leaves under the control of the constitutive 35S promoter (Figure 6). Unlike the yeast pah1Δ mutant, the volume of the nucleus was not greatly enlarged in the cells of pah1 pah2-1 leaves, and its shape was not distorted (Figures 6A and 6B). However, analysis of the ER using both lumen (HDEL) and membrane (calnexin) markers suggested that the morphology was dramatically altered (Figures 6C to 6J). The ER is usually connected to the nuclear envelope in plant cells and comprises sheet-like regions (cisternae) and an extensive cortical network of tubules, predominantly connected by three-way junctions (Staehelin, 1997). In pah1 pah2-1 cells, it appears that the network of tubules, which usually makes up the majority of the ER, is partially replaced by ER sheets (Figures 6C to 6J). This change in subcellular morphology would be consistent with the overproduction of ER-derived phospholipids and consequent expansion of the ER. Recent studies performed on the Caenorhabditis elegans Lipin homolog LPIN-1 report that downregulation of this gene also leads to the appearance of large membrane sheets and other abnormal structures in the peripheral ER (Golden et al., 2009; Gorjánácz and Mattaj, 2009). In contrast with the ER, visualization of chloroplasts via chlorophyll autofluorescence suggested that their morphology appeared relatively normal (Figures 6K and 6L).

Figure 6.
Figure 6.

Nuclear, ER, and Chloroplast Morphology in pah1 pah2-1 Double Mutant Plants.

Single plane confocal images of wild-type and pah1 pah2-1 leaf cells. Bars = 10 μm.

(A) The wild type expressing nuclear-targeted GFP.

(B) pah1 pah2-1 expressing nuclear targeted GFP.

(C) to (E) The wild type expressing ER lumen–targeted GFP.

(F) to (H) pah1 pah2-1 expressing ER lumen–targeted GFP.

(I) The wild type expressing ER membrane–targeted GFP.

(J) pah1 pah2-1 expressing ER membrane–targeted GFP.

(K) and (L) Chlorophyll autofluorescence in the wild type (K) and chlorophyll autofluorescence in pah1 pah2-1 (L). Biolistics was performed on the abaxial surface of rosette leaves.

The Net Rate of PC Synthesis Is Increased in pah1 pah2 Leaves

PC is the predominant phospholipid class in Arabidopsis tissues (Ohlrogge and Browse, 1995), and its de novo synthesis proceeds by the branched pathway depicted in Figure 7 (Keogh et al., 2009). To measure the net rate of PC synthesis via the nucleotide pathway in wild-type and pah1 pah2-1 leaves (Figure 8), radiolabel feeding experiments were performed using [methyl-14C]choline chloride as a substrate (Kinney et al., 1987; Tasseva et al., 2004). Incorporation of [methyl-14C]choline into PC was linear with respect to time for 3 h, and the rate was 1.8-fold greater in pah1 pah2-1 leaves than in the wild type, on a per unit FW basis (Figure 8A). These data suggest that the net rate of PC synthesis, via the nucleotide pathway, is substantially increased in pah1 pah2-1 leaves. The stimulation of PC synthesis may be the consequence of upregulation of specific steps of the nucleotide pathway. To investigate this possibility, changes in the proportion of radiolabel in each of the pathway intermediates was monitored following the supply of a pulse of [methyl-14C]choline chloride (Tasseva et al., 2004). Over the course of 2 h following the pulse, pah1 pah2-1 leaves incorporated a greater proportion of the radiolabel in PC than did the wild type, and a lesser proportion was observed in the phosphocholine pool (Figures 8B and 8C). Less than 3% of the radiolabel was detected in the form of CDP-choline in either wild-type or pah1 pah2-1 leaves (Figures 8B and 8C), and <1% was found in lyso-PC.

Figure 7.
Figure 7.

A Simplified Diagram to Illustrate the Pathways for PC Synthesis in Eukaryotes.

Left to right: the methylation pathway(s). Bottom to top: the nucleotide pathway(s). Steps common to plants, mammals, and yeast are indicated by black arrows. Dashed arrows indicate pathways specific to some plants, such as Arabidopsis. Methylation of PE, which occurs in mammals and yeast, is indicated by a dotted arrow. Cho, choline; EA, ethanolamine; P-, phospho; CDP, cytidine diphosphate; MM, monomethyl; DM, dimethyl; AAPT, aminoalcoholphosphotransferase; CCT, CTP-phosphocholine cytidylyltransferase; EKI, ethanolamine kinase; PAPH phosphatidic acid phosphohydrolase.

Figure 8.
Figure 8.

Incorporation of [methyl-14C]Choline into PC in pah1 pah2-1 Leaves.

(A) A time course of [methyl-14C]choline incorporation into PC. Values are means ± se of measurements from five separate incubations. WT, wild type.

(B) and (C) A time course of in vivo label distribution among water-soluble derivatives choline (Cho), phosphocholine (P-Cho), and cytidine diphosphocholine (CDP-Cho) and PC in wild-type (B) and pah1 pah2-1 (C) leaves, following a 10-min treatment with [methyl-14C]choline. Results are means ± se of measurements from three separate experiments.

Two inferences can be drawn from these data. First, the observation that the phosphocholine pool contains a far greater proportion of the radiolabel than the CDP-choline pool, in both wild-type and pah1 pah2-1 leaves, suggests that cytidine triphosphate:phosphocholine cytidylyltransferase (CCT) activity might exert a relatively high level of control over the rate of flux of choline to PC (Kinney et al., 1987; Tasseva et al., 2004). Second, the decrease in the relative proportion of radiolabel in the phosphocholine pool in pah1 pah2-1 leaves, versus the wild type, suggests that CCT activity could be increased, although it does not indicate whether or not choline kinase (CKI) activity might also be affected (Tasseva et al., 2004). To determine directly whether the activity of CCT is altered, enzyme assays were performed using crude leaf extracts (Inatsugi et al., 2002). Total CCT activity in leaves from pah1 pah2-1 plants was more than twofold greater than the wild type (0.51 ± 0.12 μmol g−1 FW min−1, verses 0.23 ± 0.05 μmol g−1 FW min−1, n = 5, ±se).

The Expression of Several Genes Involved in Phospholipid Synthesis Is Increased in pah1 pah2 Leaves

To investigate whether disruption of Arabidopsis PAH1 and PAH2 affects the expression of genes associated with phospholipid synthesis, real-time PCR was used to measure their relative transcript abundance. A survey of 13 genes revealed that several are significantly upregulated in pah1 pah2-1 leaves (Table 4). The most strongly induced of these genes is phosphoethanolamine N-methyltransferase1 (PEAMT1/XIPOTL), for which the mRNA is approximately sevenfold more abundant (Table 4). PEAMT catalyzes the first committed step of choline synthesis in many plants (Figure 7), and it is believed to be vital for PC synthesis in Arabidopsis (Bolognese and McGraw, 2000; McNeil et al., 2000, 2001; Mou et al., 2002; Cruz-Ramírez et al., 2004; Keogh et al., 2009). In addition to PEAMT1, the transcript abundance of CCT1, cytidine triphosphate:phosphoethanolamine cytidylyltransferase (ECT), and a putative choline kinase (At1g74320) are also upregulated more than twofold in pah1 pah2-1 leaves (Table 4). Smaller increases were also observed in the transcript abundance of the aminoalcoholphosphotransferase AAPT2 and the phospholipid N-methyltransferase (PLMT) (Table 4). All these genes encode enzymes that could potentially play a role in regulating the rate of PC and PE synthesis in Arabidopsis (Kinney et al., 1987; Tasseva et al., 2004; Mizoi et al., 2006; Keogh et al., 2009).

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Table 4.

Analysis of Gene Expression for Genes Associated with Phospholipid Synthesis in pah1 pah2-1 Leaves

PEAMT1 Is Induced at the Level of Transcription in pah1 pah2 Leaves

Because PEAMT1 is induced in pah1 pah2-1 leaves and defines a variant pathway for PC synthesis that is not found in yeast or mammals (Keogh et al., 2009), we chose to study its regulation in more detail. Interestingly, it has recently been reported that PEAMT1 is regulated at the posttranscriptional level via an upstream open reading frame (uORF) in its 5′ untranslated region (UTR) and that this uORF is necessary for repression of protein (and transcript) abundance by exogenous choline (Tabuchi et al., 2006). To investigate whether the induction of PEAMT1 in pah1 pah2-1 leaves is achieved by transcriptional or posttranscriptional/translational regulation, promoter-β-glucuronidase (GUS) reporter experiments were performed using two constructs containing ~1.3 kb of the PEAMT1 promoter including the 5′UTR. In one construct, the 5′UTR is intact, and in the other, a mutation has been introduced into the initiation codon of the uORF. The data presented in Figure 9 show that the PEAMT1 promoter is sufficient to drive an approximately eightfold induction of GUS activity in pah1 pah2-1 leaves, relative to the wild type, and that this induction occurs independently of the mutated uORF. Furthermore, in the presence of 10 mM choline chloride, a functional uORF is required for repression of PEAMT1 promoter-GUS activity (Figure 9). These data suggest that PEAMT1 is subject to transcriptional regulation by PAH1/2 and posttranscriptional/translational regulation by choline (or a related metabolite).

Figure 9.
Figure 9.

Promoter-Reporter Analysis of PEAMT1 Expression in Wild-Type versus pah1 pah2-1 Leaves.

(A) Diagram of the PEAMT1promoter:GUS reporter constructs showing the uORF in black. In one construct, the ATG of the uORF is intact (Plus ORF) and in the other it has been mutated to TTG (Minus ORF).

(B) Relative expression in pah1 pah2-1 leaves versus the wild type (WT) for the Plus ORF and Minus ORF constructs was determined as the difference in the ratio of PEAMT1promoter:GUS to 35S:LUC activity. Values are the mean ± se from three independent bombardment experiments, each using six leaves per construct and treatment. Leaves were either incubated with half-strength MS salts alone (control) or plus 10 mM NaCl or 10 mM choline chloride (Cho-Cl).

DISCUSSION

In this study, we show that disruption of two Mg2+-dependent PA phosphohydrolases (called PAH1 and PAH2) causes derepression of phospholipid biosynthesis and ER membrane biogenesis in Arabidopsis. Nakamura et al. (2009) also recently characterized PAH1/2 and proposed that they play a role in the eukaryotic pathway for galactolipid synthesis. This conclusion is based primarily on the observation that pah1 pah2-1 leaves exhibit a relative decrease in MGD and DGD content, on a mol % basis, and that MGD is enriched in 16:3, which is a fatty acid derived from the prokaryotic pathway (Nakamura et al., 2009). Our own analysis of pah1 pah2-1 supports these findings but also shows that the leaves exhibit an absolute increase in phospholipid content, on a FW basis, that is disproportionate to the reduction in galactolipids and results in an overall increase in leaf polar lipid content. Furthermore, an equivalent increase in phospholipid content is also seen in roots, which contain comparatively little galactolipid. It is unlikely that this effect can be interpreted solely as the result of a direct block in eukaryotic flux to galactolipids causing phospholipid precursors to backup in the ER (Nakamura et al., 2009). Instead, the data indicate that phospholipid synthesis is enhanced in pah1 pah2-1.

We cannot exclude a direct role for PAH1/2 in the provision of substrate for galactolipid synthesis, as proposed by Nakamura et al. (2009). However, it is possible to present an alternative explanation for the negative impact of pah1 pah2-1 on leaf galactolipid content, whereby increased production of phospholipids diverts flux away from the eukaryotic pathway. Nakamura et al. (2009) also proposed a role of PAH1/2 in membrane lipid remodeling during phosphate (P) starvation, when extraplastidial phospholipids are replaced by galactolipids. However, the impairment of galactolipid synthesis that Nakamura et al. (2009) report in P-starved pah1 pah2-1 plants could also be attributed to enhanced phospholipid synthesis and does not necessarily imply a physiological role of PAH1/2 in the remodeling process. Further work will be required to resolve these questions.

As described in the introduction, Pah1p also acts indirectly as a repressor of phospholipid synthesis in yeast (Carman and Henry, 2007). It is logical to speculate that the underlying mechanisms might be conserved. However, the signal transduction pathway, which couples PAH to the regulation of phospholipid synthesis in Arabidopsis, is likely to be substantially different from that described in yeast for two reasons. First, sequence similarity searches using BLAST suggest that Arabidopsis lacks homologs of the core regulatory genes, namely, the repressor OPI1 and the transcriptional regulators INO2 and INO4. These genes are instrumental for Pah1p-mediated repression of phospholipid biosynthetic gene expression in yeast (Carman and Henry, 2007). Second, there are fundamental differences between the phospholipid biosynthetic pathways of Arabidopsis and yeast (Figure 7), which appear to be reflected in the different enzymes that are regulatory targets in the two biological systems.

PC can be formed through two distinct pathways in yeast and mammals, namely, the nucleotide pathway, in which free choline is incorporated into PC using CDP-choline as an intermediate, and the methylation pathway whereby PC is produced directly from PE via three sequential methylation reactions, using S-adenosylmethionine as the methyl donor (Carman and Henry, 2007; Carman and Han, 2009; Sugimoto et al., 2008). By contrast, net PC biosynthesis in plants occurs through a branched pathway that uses components of both the nucleotide and methylation pathways (Keogh et al., 2009). The key difference can be attributed to the presence of the plant enzyme PEAMT, which is capable of converting ethanolamine head groups to choline at the phospho-base level, an activity that is absent in yeast and mammals (Bolognese and McGraw, 2000; McNeil et al., 2000, 2001; Mou et al., 2002; Cruz-Ramírez et al., 2004). Conversely, mammals and yeast possess methylation enzymes that act directly on PE, a reaction that cannot be detected in most plants that have been investigated and appears to be absent in Arabidopsis (Keogh et al., 2009).

Our data show that PEAMT1 is a target for PAH1/2-mediated repression in Arabidopsis and that this regulation is likely to occur at the level of transcription. Interestingly, it has recently been reported that PEAMT1 contains a uORF in its 5′UTR and that this uORF is necessary for repression of protein (and transcript) abundance by exogenous choline (Tabuchi et al., 2006). PEAMT1 is therefore subject to multiple layers of regulation. PEAMT is necessary for PC synthesis in Arabidopsis (Cruz-Ramírez et al., 2004) but, despite catalyzing the first committed step in the pathway, it is unclear whether it exerts a high degree of control over the net rate of synthesis. Overexpression of PEAMT in tobacco (Nicotiana tabacum) plants has been reported to increase choline but not PC content (McNeil et al., 2001). However, kinetic metabolic flux analysis performed by McNeil et al. (2000) also suggested that choline synthesis proceeds primarily via PC turnover in tobacco leaves, rather than by dephosphorylation of phosphocholine. If this is the case, PEAMT could exert significant control over the rate of metabolic flux to PC, without PC content necessarily increasing. The flux model created by McNeil et al. (2000) indicated that PC is turned over by phospholipase C and D activities in tobacco leaves, yielding phosphocholine and choline, respectively. We cannot discount the possibility that downregulation of PC turnover, as well as upregulation of synthesis, contributes to its overaccumulation in pah1 pah2-1 plants.

In addition to PEAMT1, the genes CCT1 and ECT are also repressed by PAH1 and PAH2 in Arabidopsis. By contrast, the homologous genes in yeast (PCT1 and ECT1) are unlikely to be regulated by Pah1p since they do not contain UASINO elements in their promoters (Carman and Henry, 2007), and they are not repressed by inositol (Jesch et al., 2005). CCT1 and ECT encode nucleotide pathway enzymes that are believed to play an important role in regulating the rate of PC and PE synthesis, respectively, in Arabidopsis (Kinney et al., 1987; Tasseva et al., 2004; Mizoi et al., 2006). Data from our own radiolabel feeding experiments also suggest that the supply of CDP-choline is most likely to be rate-limiting for PC synthesis in Arabidopsis leaves and that this supply is increased in the pah1 pah2-1 mutant by an enhanced flux through CCT, whose in vitro activity in a crude leaf extract also increases more than twofold. Finally, CKI-like2 (a putative choline kinase gene), AAPT2, and PLMT are also repressed, to varying degrees, by PAH1 and PAH2 in Arabidopsis. Homologous genes in yeast (CKI1, CPT1, and OPI3) are likely to be regulated by Pah1p (Jesch et al., 2005; Carman and Henry, 2007).

Based on catalytic function alone, it is perhaps counterintuitive that a reduction in PAH1/2 activity would lead to an increase in PC and PE levels in plants, since the product of the reaction (diacylglycerol [DG]) is required for their synthesis via the nucleotide pathway (Figure 7). However, plants contain multiple PA phosphohydrolase isozymes (Pierrugues et al., 2001; Nakamura et al., 2007), and pah1 pah2-1 plants still contain a substantial amount of PA phosphohydrolase activity (Nakamura et al., 2009). Radiolabel feeding experiments also suggest that the supply of choline head groups, rather than DG, is most likely to be rate-limiting for PC synthesis (Kinney et al., 1987; Tasseva et al., 2004). In the majority of mammalian tissues, PC is also synthesized by the nucleotide pathway, and CCT is well established as the key regulatory enzyme in this pathway (Sugimoto et al., 2008). CCT associates with the ER membrane surface via an amphipathic helix. The protein only becomes active when it translocates to the ER and its activity is modulated by the composition (and curvature) of the membrane. High levels of DG, PA, and free fatty acids stimulate CCT activity (Sugimoto et al., 2008). In plants, the importance of CCT posttranslational modification or allosteric regulation is not clear. Although plant CCT activity can be modulated by lipids, the effects do not generally appear to be as large as have been reported in mammals (Price-Jones and Harwood, 1983, 1986; Kinney and Moore, 1987). Nevertheless, posttranslational modification or allosteric regulation of enzymes such as CCT could play a significant role in the modulation of phospholipid synthesis by PAH1/2.

It remains to be determined whether the repression of phospholipid synthesis in Arabidopsis is mediated directly by the PAH proteins or by the level of a key lipid metabolite, which they regulate, such as PA. The pah1 pah2-1 mutant does accumulate PA in its leaves, and this could potentially trigger the induction of phospholipid synthesis. PA is already known to be an important phospholipid signal in plants (Munnik, 2001; Wang et al., 2006). Catalytically dead forms of Pah1p are unable to complement any of the pah1Δ mutant’s phenotypes, supporting a specific signaling role for PA in the regulation of phospholipid synthesis in yeast (Han et al., 2007). A yeast DG kinase mutant (dgk1Δ) has also recently been characterized, which suppresses pah1Δ, while overexpression of DGK1 phenocopies pah1Δ (Han et al., 2008). There is indirect evidence to suggest that Lipins might also be involved in the regulation of phospholipid synthesis in animals (Kim et al., 2007). In C. elegans, the downregulation of LPIN-1 leads to changes in ER morphology that are broadly similar to those we observed in Arabidopsis pah1 pah2-1 (Golden et al., 2009; Gorjánácz and Mattaj, 2009). However, whether this effect in C. elegans is mediated by PA is unknown at present. Alternatively, Finck et al. (2006) reported that Lipin-1 has a transcriptional role in mammals that is independent of its catalytic activity. Lipin-1 apparently functions as a transcriptional coactivator by interacting directly with the long-chain fatty acid receptor/transcription factor PPARα, leading to the formation of a complex that could potentially modulate the expression of multiple genes involved in lipid metabolism (Finck et al., 2006; Donkor et al., 2009). For Arabidopsis PAH1/2 to function in this way, they would need to locate to the nucleus. However, neither protein was found in the nucleus when expressed as N-terminal GFP fusions.

Yeast Pah1p and mammalian Lipins play an important structural role in TG synthesis, catalyzing the penultimate step in the pathway. The yeast pah1Δ mutant is severely compromised in TG synthesis during the stationary phase of the cell cycle when TG predominantly accumulates (Han et al., 2006). In mice, a defect in Lipin-1 also causes a reduction in adipose tissue mass (Péterfy et al., 2001). It is surprising therefore that in the Arabidopsis pah1 pah2-1 double mutant, seed TG content is not substantially reduced relative to the wild type. Interestingly, a recent study using developing soybean (Glycine max) embryos suggests that de novo synthesis of DG is mainly used for PC synthesis, whereas another kinetically distinct DG pool that is mostly derived from PC turnover supports TG synthesis (Bates et al., 2009). On this basis, PAH need not catalyze the penultimate step in TG biosynthesis. Nevertheless, DG production must keep pace with the overall demand for glycerolipids during both the vegetative and reproductive phases of pah1 pah2-1 development. The simplest explanation is that, in the absence of PAH1/2, other PA phosphohydrolase isozymes (Pearce and Slabas, 1998; Pierrugues et al., 2001; Nakamura et al., 2007, 2009) enable this to happen. In wild-type plants, PAH1/2 may still play a predominant role in eukaryotic glycerolipid synthesis. Both genes are expressed most strongly during late seed development, when the demand for glycerolipids is greatest, and the proteins specifically catalyze the dephosphorylation of PA and not lyso-PA in vitro. This substrate selectivity has not been shown previously among the plant PA phosphohydrolases that have been characterized, and it would be consistent with a specific metabolic role within the Kennedy pathway (Pearce and Slabas, 1998; Pierrugues et al., 2001; Nakamura et al., 2007).

In conclusion, our analysis of the pah1 pah2-1 mutant suggests that PAH1/2 have both metabolic and signaling roles in Arabidopsis and that the hypothesis that they play a direct role in the provision of substrate for the eukaryotic pathway of galactolipid synthesis (Nakamura et al., 2009) may need to be reevaluated. The pah1 pah2-1 mutant exhibits an absolute increase in phospholipid content that is disproportionate to the decrease in galactolipids. The net rate of PC synthesis, the expression of several phospholipid biosynthetic genes, and ER morphology are all altered. These data suggest that PAH1/2 play a role in repressing phospholipid synthesis at the ER. The signal transduction mechanism by which PAH1/2 regulates phospholipid synthesis in Arabidopsis remains unknown and will require further study. By analogy with mammalian and yeast systems (Finck et al., 2006; Carman and Henry 2007), PAH1/2 may either be bifunctional proteins capable of modulating phospholipid synthesis independently of their catalytic activity or they might govern phospholipid synthesis through changes in the level of a key metabolite, such as PA or DG.

METHODS

Plant Material and Growth Conditions

Wild-type Arabidopsis thaliana ecotype Columbia-0 and the pah1 and pah2-1 mutants (SALK_042970 and SALK_047457; Alonso et al., 2003) were obtained from the European Arabidopsis Stock Centre (University of Nottingham, UK). The pah2-2 (FLAG_076D10) and pah2-3 (FLAG_196D12) mutants were provided by INRA (Brunaud et al., 2002). Seeds were surface sterilized, applied to agar plates containing half-strength MS salts (Sigma-Aldrich), and imbibed in the dark for 4 d at 4°C. The plates were then transferred to a growth chamber set to 21°C 16 h light/18°C 8 h dark; PPFD = 150 μmol m−2 s−1. Seedlings were transferred from plates to soil after 7 d and grown to seed under the same conditions as described above.

Cloning, Expression, and Purification of PAH1 and PAH2

For expression of PA phosphohydrolases in Escherichia coli, the entire coding sequence of the two genes was amplified from cDNA by PCR using the primer pairs 5′-CACCGCTAGCGTTGGAAGAGTTGGGAGTTTGATTTCTCAAGG-3′ and 5′-CACCCTCGAGTCATTCAACCTCTTCTATTGGCAGTTTCC-3′ for PAH1 and 5′-CACCGCTAGCGTCGGTAGGATCGGAAGCTATATCTACC-3′ and 5′-CACCCTCGAGTCACATAAGCGATGGAGGAGGCAGTTTCCAG-3′ for PAH2. The PCR products were then cloned into the pET-28a expression vector using the NheI/XhoI sites and introduced into the E. coli strain ArcticExpress RIL (Stratagene). E. coli cells were grown at 37°C in Luria-Bertani medium, containing 50 μg/mL kanamycin and 25 μg/mL gentamycin. A new Luria-Bertani culture was inoculated from the overnight culture (1:50, v/v) and incubated at 30°C for 3 h, followed by temperature equilibration at 13°C for ~30 min. The culture was then incubated for 20 h with 1 mM isopropyl β-d-thiogalactopyranoside to induce the expression of the His6-tagged recombinant proteins.

All steps for protein purification were performed at 4°C. E. coli cells expressing His6-tagged recombinant proteins were washed once with 20 mM Tris-HCl buffer, pH 8.0, and suspended in 40 mL of Buffer A: 20 mM sodium phosphate, pH 7.7, 0.5 M NaCl, 20 mM imidazole, 7 mM 2-mercaptoethanol, and 1:20 (v/v) Protease Inhibitor Cocktail (Sigma-Aldrich). Cells were disrupted by resuspension with a glass homogenizer followed by three passes through a French press at 25,000 p.s.i. Unbroken cells and cell debris were removed by centrifugation at 40,000g for 30 min at 4°C. The supernatant (cell lysate) was filtered through a 20-μm filter and the sample loaded on a 5 mL Ni2-NTA column (Qiagen) followed by a 5-mL wash with Buffer A. His6-tagged proteins were then eluted from the column in 1-mL fractions with a total of 50 mL of a linear gradient of Buffer A containing up to 500 mM imidazole. The fractions were analyzed via 7.5% SDS-PAGE and stained with Coommassie Brilliant Blue. Pooled enzyme preparations were dialyzed against 20 mM Tris-HCl buffer, pH 7.0, containing 10% (v/v) glycerol and 7 mM 2-mercaptoethanol and stored at −80°C.

Radiolabel Assays

[32P]PA was synthesized enzymatically from DG and [γ-32P]ATP using E. coli DG kinase (Calbiochem) in a total reaction volume of 150 μL. The final concentration of the reagents was 51 mM (15 mg/mL) octylglucoside, 1 mM cardiolipin, 60 mM imidazole-HCl, pH 6.6, 50 mM NaCl, 12.5 mM MgCl2, 1 mM EGTA, 0.3 mM DPTA, 2 mM DTT, 2 mM dioleoylglycerol, 50 μM [γ-32P]ATP (30 Ci/mmol, 10 mCi/mL, and 150 μCi total), and 0.05 units of DG kinase. The reaction was incubated at 25°C for 1 h followed by termination with 90 μL of 10% perchloric acid. Phase extraction was performed with 2.5 mL of chloroform/methanol (1:1) and 1 mL of 0.2 M H3PO4 in 1 M KCl. The organic phase was reextracted (washed) with 1 mL of 0.2 M H3PO4 in 1 M KCl, and the aqueous phase was reextracted with 2.5 mL of chloroform/methanol (1:1). The two organic phases were then pooled and washed once more with 1 mL of 0.2 M H3PO4 in 1 M KCl, and 1 μL was subjected to liquid scintillation counting, typically yielding ~100,000 dpm. The purity of the [32P]PA was confirmed by TLC using chloroform/methanol/water (12:6:1 by volume) as the developer.

Mg2+-dependent PA hydrolase activity was measured by following the release of water-soluble 32Pi from chloroform-soluble [32P]PA (10,000 cpm/nmol) by a modification of the protocol described by Carman and Lin (1991). The standard reaction mixture contained 50 mM Tris-HCl buffer, pH 6.5, 1 mM MgCl2, 0.3 mM PA, 3 mM Triton X-100, and enzyme protein in a total volume of 0.1 mL. All enzyme assays were conducted in triplicate at 30°C. The reaction was terminated by removing 100 μL into 720 μL chloroform/methanol (1:1) and 280 μL 1 M KCl/0.2 M H3PO4, and after mixing, phase separation was achieved by centrifugation at 4000 rpm for 5 min. Ecoscint A (National Diagnostics) was added to 100 μL of aqueous phase, and liquid scintillation counting was used to determine the radioactivity.

For radiolabel feeding experiments, rosette leaves (0.2 g FW) were cut into fragments using a razor blade and placed in vials containing 0.5 mL of half-strength MS salts supplemented with 0.5 μCi/mL [methyl-14C]choline chloride (56 mCi/mmol; Sigma-Aldrich). The leaf tissue was vacuum infiltrated for 5 min. For measurements of the proportion of radiolabel in all choline-containing intermediates, the tissue was then rinsed with water to remove excess choline chloride and resuspended in half-strength MS. Following incubation at 22°C with gentle agitation, the tissue was gently blotted dry and frozen in liquid nitrogen. The extraction and analysis of labeled choline-containing metabolites was conducted according to the methods described by Tasseva et al. (2004). CCT assays were performed on crude leaf extracts at 25°C following the method described by Inatsugi et al. (2002). The rate of the reaction was confirmed to be linear with time for up to 30 min and also to be proportional to the amount of extract included in the assay.

Lipid Analysis

The fatty acid composition of seeds, roots, and leaf tissue was measured by GC analysis after combined digestion and fatty acid methyl ester formation using the method of Browse et al. (1986). The TG content of seeds was determined using the method described by Eastmond (2006). Lipids were extracted from 0.2 g FW of leaf tissue according to Dörmann et al. (1995). The extracts were applied to silica gel 60A plates (Merck) and analyzed by two-dimensional TLC, using chloroform/methanol/water (75:25:2.5, by volume), followed by chloroform/methanol/acetic acid/water (80:9:12:2, by volume) according to the method of Christie (2003). Lipids were visualized under UV light having been sprayed with primulin solution (0.01% [w/v] in 60% [v/v] acetone). The lipids were identified by cochromatography with commercial standards (Sigma-Aldrich) and their fatty acid content determined by GC analysis (Browse et al., 1986). To allow quantification, the lipid extracts were spiked with 15:0 TG, and the absolute quantity of each lipid class was calculated based on the final recovery of 15:0. Positional analysis of fatty acids in PA and separation of TG and TGD by TLC were done as described by Xu et al. (2005). PC was quantified directly from crude tissue extracts using the enzyme-coupled fluorometric procedure described by Nanjee et al. (1991). The recovery of PC was determined to be between 95 and 106% using this method.

Photosynthetic Parameters

Gas exchange measurements were made using a CIRAS-2 portable photosynthesis system with a PLCAR leaf cuvette (PP Systems). Chlorophyll fluorescence was determined using a cfImager (Technologia Ltd.). Chlorophyll content was measured using the method of Arnon (1949) with modifications described by Porra (2002).

Transcript Analysis

DNase-treated total RNA was isolated from Arabidopsis tissues using the RNeasy kit from Qiagen . In the case of seeds, this method was modified as described by Wu et al. (2002). The synthesis of single-stranded cDNA was performed using SuperScript II RNase H- reverse transcriptase from Invitrogen. PAH1 and PAH2 transcripts were detected in wild-type, pah1, and pah2-1 plants by PCR using primer pairs for PAH1 (5′-AAATGAGTTTGGTTGGAAGAGTTGGGAG-3′ and 5′-AGCTCCCAAGAAGATCTCTTTCAGCATC-3′) and PAH2 (5′-CCAAGGGCATGATAGCTGTT-3′ and 5′-TCACATAAGCGATGGAGGAG-3′). ACT2 primers are described by Eastmond et al. (2002).

Quantitative real-time PCR was performed with an ICycler (Bio-Rad) using qPCR Mastermix Plus (Eurogentec), according to the manufacturer’s instructions, and the data were analyzed with the ICycler IQ5 software. To calculate the PCR efficiency, serial dilutions of cDNA were used as template in the presence of the different primer pairs and the cycle threshold (CT) was plotted against the initial quantities of reverse-transcribed RNA. The slope of the resulting curve allowed the calculation of PCR efficiency (E) according to the equation E = 10[−1/slope]. Primer sequences and PCR efficiencies are given in Supplemental Table 2 online. The relative expression ratio (R) of the target genes was calculated based on E and the deviation in the number of cycles required to reach threshold values (ΔCT) in comparison to a reference gene (Pfaffl, 2001). ACTIN2/ACTIN8 were chosen as the reference gene for this study as their expression has been shown to be relatively invariant (An et al., 1996). However, in control experiments, it was confirmed that primers for 18S (Eastmond, 2006) yielded similar results.

Transient Expression Experiments

The ER lumen marker 35S:GFP-HDEL and the nuclear marker 35S:GFP-GFP-NLS were obtained from Katrin Czempinski and Bernd Müller-Röber (University of Potsdam, Germany). The ER membrane marker 35S:GFP-calnexin (Irons et al., 2003) was provided by Lorenzo Frigerio (University of Warwick). Transfer into the abaxial epidermal cells of Arabidopsis leaves was achieved by bombardment in a Biolistic PSD-1000/He particle delivery system (Bio-Rad) using 1100 p.s.i. rupture discs according to the manufacturer's protocol. Plasmid DNA (2.5 μg) was coated onto 3 mg of 1.2-μm tungsten particles (Bio-Rad). Leaves were incubated for 48 h in low light at 20°C with their cut petiole inserted into half-strength strength MS salts solution. Transient expression in Nicotiana benthamiana leaf epidermal cells was performed by agroinfiltration as described by Sparkes et al. (2006), using agrobacteria at an OD600 of 0.1. Fluorescence signals were analyzed using a Leica TCS SP2 or a Zeiss LSM 710 confocal microscope.

The PEAMT1 promoter was amplified from wild-type genomic DNA using the primers PEAMT1P5 (5′-CACCGTCGTGGTGGTGGATATGTAAATGTG-3′) and PEAMT1P3 (5′-TTTCGGAAATGTCGTTTGTCG-3′). A point mutation was introduced into the PEAMT1 promoter by PCR using PEAMT1P5 and PEAMT1P3 in combination with 5′-TCTCTGCTGCAACAATTCGCACAACGTC-3′ and 5′-GACGTTGTGCGAATTGTTGCAGCAGAGA-3′, according to the method described by Ho et al. (1989). The products were cloned using the pENTRY directional TOPO cloning kit from Invitrogen and then transferred into the plant expression vector pBGWFS7 (Karimi et al., 2002) by recombination, using the LR clonase enzyme mix (Invitrogen). Bombardment experiments were performed as described above, except that PEAMT1promoter:GUS plasmids were cobombarded with 35S:Luciferase (Luc) at a ratio of 3:1, and 10 mM choline chloride or NaCl was included in the half-strength MS salts solution. Leaves were homogenized in 1 mL of extraction buffer (100 mM potassium phosphate, pH 7.8, 2 mM DTT, 2 mM EDTA, and 5% glycerol). The homogenate was centrifuged at 15,000g for 30 min at 4°C, and the GUS and Luc activities were assayed in the supernatant using the methods described by Lanahan et al. (1992). The values for GUS activity were normalized with respect to Luc.

Accession Numbers

Sequence data from this article can be found in the Arabidopsis Genome Initiative or GenBank/EMBL databases under the following accession numbers: At3g09560 or FJ591133 for PAH1 and At5g42870 or FJ591134 for PAH2.

Author Contributions

P.J.E. and A.R.S. conceived and designed the experiments. P.J.E., A.-L.Q., J.T.M.K., C.C., and N.A. performed the experiments and analyzed the data. P.J.E. wrote the article.

Supplemental Data

The following materials are available in the online version of this article.

Acknowledgments

We thank Lorenzo Frigerio (University of Warwick) for his invaluable assistance with confocal microscopy. The work performed by A.-L.Q., C.C., N.A., and P.J.E. was funded by the Biotechnology and Biological Sciences Research Council (BBSRC). J.T.M.K. and A.R.S. were supported by a cooperative award jointly funded by the BBSRC and Arkema. A.R.S. thanks Linnaeus Plant Sciences for its continual support.

Footnotes

  • The author responsible for distribution of materials integral to the findings presented in this article in accordance with the policy described in the Instructions for Authors (www.plantcell.org) is: Peter J. Eastmond (p.j.eastmond{at}warwick.ac.uk).

  • www.plantcell.org/cgi/doi/10.1105/tpc.109.071423

  • ↵1 These authors contributed equally to this work.

  • ↵2 Current address: Institut des Sciences du Végétal, CNRS-UPR2355, 91198 Gif-sur-Yvette cedex, France.

  • ↵[W] The online version of this article contains Web-only data.

  • Received September 25, 2009.
  • Revised June 22, 2010.
  • Accepted July 21, 2010.
  • Published August 10, 2010.

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PHOSPHATIDIC ACID PHOSPHOHYDROLASE1 and 2 Regulate Phospholipid Synthesis at the Endoplasmic Reticulum in Arabidopsis
Peter J. Eastmond, Anne-Laure Quettier, Johan T.M. Kroon, Christian Craddock, Nicolette Adams, Antoni R. Slabas
The Plant Cell Aug 2010, 22 (8) 2796-2811; DOI: 10.1105/tpc.109.071423
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