Involvement of phosphatidate phosphatase in the biosynthesis of triacylglycerols in Chlamydomonas reinhardtii

Abstract

Lipid biosynthesis is essential for eukaryotic cells, but the mechanisms of the process in microalgae remain poorly understood. Phosphatidic acid phosphohydrolase or 3-sn-phosphatidate phosphohydrolase (PAP) catalyzes the dephosphorylation of phosphatidic acid to form diacylglycerols and inorganic orthophosphates. This reaction is integral in the synthesis of triacylglycerols. In this study, the mRNA level of the PAP isoform CrPAP2 in a species of Chlamydomonas was found to increase in nitrogen-free conditions. Silencing of the CrPAP2 gene using RNA interference resulted in the decline of lipid content by 2.4%–17.4%. By contrast, over-expression of the CrPAP2 gene resulted in an increase in lipid content by 7.5%–21.8%. These observations indicate that regulation of the CrPAP2 gene can control the lipid content of the algal cells. In vitro CrPAP2 enzyme activity assay indicated that the cloned CrPAP2 gene exhibited biological activities.

1. Introduction

With rapidly decreasing fossil fuel resources, the importance of energy and environmental preservation has received increasing interest. Microalgae biodiesel, a crucial part of renewable biomass energy that uses solar energy to convert CO₂ into biomass, is the most promising alternative for fossil fuels. However, the study of lipid metabolism in the eukaryotic single-celled photosynthetic microalgae has lagged behind that of oil crops. Basic knowledge of microalgae has long fallen behind that of most crops, such as rice, rape, and corn. However, with the increasing number of microalgae-derived biodiesel studies on a global scale, more research teams are focusing on the underlying mechanisms of high lipid production and high cell density cultures. These processes are crucial to the improvement of genetic strains, and to the future cultivation of commercial and industrial microalgae. Phosphatidic acid phosphohydrolase (3-sn-phosphatidate phosphohydrolase, PAP, EC 3.1.3.4) catalyzes the dephosphorylation of phosphatidic acid to form diacylglycerols and inorganic orthophosphates (Smith et al., 1957). Depending on their requirement for Mg²⁺, PAPs are classified as either Mg²⁺-dependent or Mg²⁺-independent. An Mg²⁺-dependent PAP is referred to as PAP1, whereas an Mg²⁺-independent PAP is generally referred to as lipid phosphate phosphatase (LPP) or PAP2. PAPs are involved in lipid biosynthesis or lipid signaling (Carman, 1997; Sciorra and Morris, 2002; Nanjundan and Possmayer, 2003). In mammalian cells, PAP2 reportedly participates in lipid signal transduction (Brindley, 2004), but the function of PAP1 remains unknown.

In eukaryotic cells, triacylglycerol (TAG) biosynthesis is important. This process starts from glycerol-3-phosphate and Acyl-coenzyme A to form lysophosphatidic acid. This reaction is catalyzed by glycerol-3-phosphate acyltransferase. Then, catalysis is performed through lysophosphatidyl acyltransferase to produce phosphatidic acid, which forms diacylglycerol (DAG) in the PAP reaction (Brindley, 1984). DAG is used not only to synthesize TAG, but also in the syntheses of phosphatidylethanolamine and phosphatidylcholine, which are the main constituents of membranes (Carman and Henry, 1999; Sorger and Daum, 2003). Phosphatidic acid through CDP-diacylglycerol by CDP-diacylglycerol synthetase is an alternative pathway to synthesize membrane phospholipids and their derivatives. Moreover, PAPs can regulate phospholipid synthesis at the transcriptional level (Santos-Rosa et al., 2005). The production of DAG by PAP to activate protein kinase C is an important signal pathway in response to stress (Exton, 1994; Testerink and Munnik, 2005; Howe and McMaster, 2006). The four PAP homologous genes PAH1, DPP1, LPP1, and APP1 are responsible for the PAP activity detectable in yeasts. DPP1 and LPP1 are involved in lipid signaling; they contain six transmembrane domains and a phosphatase domain. They localize in the vacuoles and Golgi body complexes, and use phosphatidic acid (PA), diacylglycerol pyrophosphate (DGPP), lysophosphatidylcholine (LysoPA), and sphingosine 1-phosphate (S1P) as substrates. PAH1 encodes the only PAP enzyme that is essential to lipid biosynthesis in Saccharomyces cerevisiae. The actin patch protein (App1) interacts with endocytic proteins, and may be involved in vesicular transportation through its PAP activity (Chae et al., 2012). In Arabidopsis, three of the PAP homologous genes, namely, AtLPP1 (lipid phosphate phosphatase, LPP), AtLPP2, and AtLPP3, have been identified. AtLPP1 encodes a 35-kD protein with a phosphatase domain and six transmembrane domains. It has high protein sequence homology with yeast DPP1. After the expression of AtLPP1 in the S double mutant dpp1Δlpp1Δ, AtLPP1 exhibits DGPP and PA phosphatase activities (Pierrugues et al., 2001). Moreover, AtLPP1 tends to use DGPP as a substrate, whereas AtLPP2 has no substrate use tendencies. AtLPP1 is expressed mainly in the leaves and roots of Arabidopsis, whereas AtLPP2 is expressed in all parts. Unlike AtLPP2, the gene expression of AtLPP1 improves when Arabidopsis is treated with ionization radiation, ultraviolet-B (UV-B) radiation, or mast cell-degranulating peptides. Thus, we conclude that AtLPP1 has an important role in the response to abiotic stress.

Merchant et al. (2007) predicted three of the PAP-homologous genes in Chlamydomonas: PAP1, PAP2, and PAH1. Of the three genes, only PAP2 has a full-length coding sequence. Thus far, no evidence has demonstrated that CrPAP2 gene expression and regulation are related to cellular lipid accumulation in Chlamydomonas. Accordingly, this study aimed to determine whether such a relationship exists. High levels of mRNA of CrPAP2 and lipid accumulation were detected in C. reinhardtii CC124 in the presence and absence of nitrogen. Suppression by RNAi and over-expression of the CrPAP2 gene were then performed in Chlamydomonas to ascertain whether the over-expression or inhibition of CrPAP2 affected lipid accumulation.

2. Materials and methods

2.1. Bioinformatics analysis of PAP2

Information on the Chlamydomonas PAP2 gene (JGI Protein ID: 343983) was obtained from the JGI Chlamydomonas database. A transmembrane assay was conducted using TMHMM 2.0. Euk-mPLoc 2.0 was used to predict the subcellular localization of proteins (Chou and Shen, 2008; 2010a; 2010b; 2010c; Chou et al., 2011; 2012; Wu et al., 2011; Chou, 2013). Sequence alignment and the phylogenetic tree of the PAP2 were created using MEGA version 4.1 (Tamura et al., 2007). Active consensus sites were identified based on the Sanger Pfam database (http://pfam.sanger.ac.uk/search).

2.2. Cultivation conditions and biomass analysis of Chlamydomonas

Algal strain C. reinhardtii CC425 (mt) was used as the receptor strain in a transgenic assay using tris acetate phosphate (TAP) agar plates and high salt medium (HSM) liquid medium for algae cultivation (Harris, 1989; Deng et al., 2011). All algae strains used in this study are listed in Table 1. For HSM-N medium, the components were the same as HSM, except for the replacement of NaCl with NH₄Cl. The cultured cells were stored in an incubator at a light intensity of 150 μmol/(m²·s) or on a shaker at 230 r/min at 25 °C.

Table 1.

Algae strain names used in this study

Name	Strain
C. reinhardtii CC425	cw-15, arg-2
Maa7-4 (10, 19)	pMaa7IR/XIR transgenic algae strains
PAP2-RNAi-3 (10, 56)	CrPAP2 RNAi transgenic algae strains
pCAMBIA-2 (8, 16)	pCAMBIA1302 transgenic algae strains
pCAMBIA-PAP2-4 (26, 60)	CrPAP2 over-expression transgenic algae strains

The algal biomass (g/L) of samples was detected at an optical density of 490 nm (OD₄₉₀), as described in a previous study (Deng et al., 2012). To generate the standard curve of OD₄₉₀ versus biomass (g/L) and guarantee that the OD₄₉₀ values ranged from 0.15 to 0.75, a series of C. reinhardtii CC425 samples were collected and diluted to appropriate ratios. The dry cell weights and OD₄₉₀ values of samples were detected. According to the results of the standard curve, the biomass was calculated using the following formula: dry cell weight (g/L)=0.7444×OD₄₉₀−0.0132 (supplementary Fig. S1).

2.3. Analysis of algal lipid content

To determine the neutral lipid level, a fluorescence method was used in accordance with the description of Deng et al. (2011). To generate the standard curve of the neutral lipid content and the fluorescence value, different concentrations of triolein (Sigma, USA) were used to measure fluorescence values after staining with Nile Red. The following formula was used to detect the algal lipid content: lipid content (g/g)=(0.0004×FD_470/570−0.0038)×0.05/dry cell weight, where FD_470/570 is the fluorescence value with an excitation wavelength of 470 nm and an emission wavelength of 570 nm (supplementary Fig. S2). For the microscopic assay, samples were stained with 0.1 mg/ml Nile Red. Results were acquired using a fluorescence microscope (Nikon 80i) (Gao et al., 2008; Chen et al., 2009; Huang et al., 2009).

2.4. RNA extraction and cloning of CrPAP2 gene

The CrPAP2 gene was amplified by polymerase chain reaction (PCR) using total RNA prepared through a modified method of Li et al. (2012). Cells from 10 ml of cultivated algae were collected by centrifugation at 10 000×g for 1 min. The supernatant was extracted using phenol and chloroform. Total RNA was precipitated with ethanol and dissolved with RNase-free water. For reverse transcription, the cDNA of C. reinhardtii CC425 was synthesized and diluted 10 times using the template for amplifying the CrPAP2 gene. PCR reactions were performed with two primers, namely, PAP2L (5′-ATTTTAGCGTTGTCGCCACT-3′) and PAP2R (5′-AGCAGCCAATTTGGTTTTGT-3′), and templates in a final volume of 50 μl of the reaction system. The resulting DNA fragment was purified using a DNA gel extraction kit (BBI, Canada), and inserted into the cloning vector pMD18-T. The constructs were identified by restriction enzyme digestion and sequencing. The sequenced plasmid was named pMD18T-PAP2.

2.5. Generating RNAi vector against CrPAP2 gene

To generate the RNAi vector for knockdown of CrPAP2 gene, we used plasmid pMD18T-PAP2 as a template, as well as primers PAP2RNAiL (5′-GCGTGTTTGCCTACTTCCTC-3′) and PAP2RNAiR (5′-CACTACTCGCGCCGTACAT-3′), to amplify the fragment of CrPAP2 and its reverse complement sequences. Then, the amplified fragments were digested with HindIII/SalI and KpnI/BamHI, and cloned into pMD18T-18S to obtain the construct pMD18-CrPAP2F-18S-CrPAP2R, which contained the inverted repeat sequence of CrPAP2 (CrPAP2 IR). Finally, CrPAP2 IR was digested with KpnI and HindIII, and inserted into pMaa7/XIR to obtain pMaa7IR/CrPAP2 IR.

2.6. Construction of the over-expression vector of CrPAP2 gene for Chlamydomonas

To construct the over-expression vector of CrPAP2 gene, the coding sequence of CrPAP2 was amplified by PCR using pMD18T-PAP2 as a template and primers 5′-TAGTAGATCTGATGGACGCGGTGACCAC-3′ and 5′-TATAACTAGTTCACCCCGTGTTCTGCATGCC-3′. The fragment was then digested with NcoI/SpeI and inserted into a similarly-digested pCAMBIA1302 to give pCAMPAP2, which allows over-expression of CrPAP2.

2.7. Transformation of Chlamydomonas

A modified glass bead method described by Kindle (1990) was used for the transformation of C. reinhardtii strain CC425. A single colony of algae was inoculated into 50 ml of TAP medium and cultivated for about 2–3 d to reach a cell density of 2×10⁶ cells/ml. These cells were used as recipients for transformation after collection and dilution by TAP. For transformation, sterile glass beads, DNA, recipient cells, and polyethylene glycol were mixed using a vortex. Finally, the mixtures were incubated in TAP plates for 7 d until the transgenic strain appeared in the medium. For over-expression of PAP2 gene, 50 μg/ml hygromycin TAP medium was used. Knockdown of PAP2 by RNAi was performed using TAP medium containing 5 μmol/L 5-FI, 5 μg/ml paromomycin, and 1.5 mmol/L L-tryptophan.

2.8. Expression of CrPAP2 in Escherichia coli BL21

To express CrPAP2 in E. coli BL21, the coding region was amplified from pMD18T-PAP2 with the primer pairs GEXPPAP2F (5′-TATGGGATCCATGGTCAATTGGAATAGT-3′) and GEXPAP2R (5′-TATAGTCGACCTAGTATCGCCCACTGAC-3′). The amplified fragments were digested with BamHI/SalI and inserted into a similarly-digested pGEX-6p-1 to give pGEXPAP2. Transformation of E. coli BL21, subsequent foreign protein detection using sodium dodecyl sulfate polyacrylamide gel electrophoresis (SDS-PAGE), and purification of the glutathione S-transferase (GST)-fusion protein were performed as described by Sambrook and Russell (2001).

2.9. Quantitative identification of PAP2 RNA

Real-time PCR was used to analyze the RNA level of PAP2 gene in HSM or HSM-N conditions, and detect the RNA level of PAP2 in RNAi and the over-expressed transgenic line. RNA samples were prepared using TRIzol, as described by Fei and Deng (2007). After cDNA was synthesized and used as a template, real-time PCR was performed on a BioRad iCycler using the PAP2 gene-specific primers PAP2RNAiL (5′-GCGTGTTTGCCTACTTCCTC-3′) and PAP2RNAiR (5′-CACTACTCGCGCCGTACAT-3′) to calculate the quantity of PAP2 RNA. SYBR Green was used as the fluorescent dye. The 18S rRNA of Chlamydomonas was used as an internal control with primers 5′-CCGTGTCAGGATTGGGTAATTT-3′ and 5′-TCAACTTTCGATGGTAGGATAGTG-3′. All reactions were performed three times. We calculated the amplification rate using a baseline-subtraction method. Relative differences in RNA were evaluated using the 2^−ΔΔC_T method, as described by Livak and Schmittgen (2001).

2.10. Measurement of PAP activities

The method of Ullah et al. (2012) was used to measure the CrPAP2 activities. The amount of Pi in nmoles released by PAP from the substrate dioleoyl-phosphatidate (1,2-dioleoyl-sn-glycero-3-phosphate, sodium salt) was determined. The reaction was stopped by the addition of 2 ml AMA reagent (acetone: 10 mmol/L ammonium molybdate:5 mol/L sulfuric acid mixture, 2:1:1 (v/v/v)), followed by 100 ml of 1 mol/L citrate solution. The resulting turbidity was removed using centrifugation at 12 000×g for 6 min at room temperature. The absorbance of the resulting yellow color was read at 355 nm using a spectrophotometer. The enzyme unit, kat, was defined as the moles of the substrate converted per second (Heinonen and Lahti, 1981; Ullah et al., 2005).

3. Results

3.1. Cloning of CrPAP2 gene and bioinformatics analysis

The gene sequence of PAP2 listed in the JGI database (JGI Protein ID: 343983) was used for the design of primers for the amplification of the gene. A full-length CrPAP2 gene was amplified by PCR. A DNA fragment of about 1 000 bp was inserted into the pMD-18T vector and sequenced. Compared with the 343983 sequence in the JGI C. reinhardtii v4.0 database, the cloned CrPAP2 CDS had a sequence with 978 bp and showed 100% consistency. This cloned fragment was named CrPAP2.

We constructed a phylogenetic tree of the members of the lipid phosphate phosphatase protein family, including C. reinhardtii CrPAP2 and Arabidopsis AtLpp1p, AtLpp2p, and AtLpp3p proteins (Fig. 1). C. reinhardtii CrPAP2 was grouped with the putative Caenorhabditis elegans phosphate phosphatase protein Q19403 in one branch, whereas Arabidopsis AtLpp1p, AtLpp2p, and AtLpp3p proteins and S. cerevisiae Dpp1p and Lpp1p proteins were grouped in a main branch (Fig. 1). Analyses indicated that C. reinhardtii CrPAP2 was a lipid phosphate phosphatase enzyme. Moreover, the location of CrPAP2 was predicted by Euk-mPLoc 2.0 to be in the membrane because CrPAP2 has five transmembrane helices (TMHMM result). As TAG can be synthesized in the endoplasmic reticulum (ER) and in the chloroplasts of Chlamydomonas, the possible location of CrPAP2 is in the membranes of ER or chloroplasts (Liu and Benning, 2013).

Fig. 1.

Phylogenetic analysis of the lipid phosphate phosphatase protein family

C. reinhardtii CrPAP2, Arabidopsis AtLpp1p, AtLpp2p, and AtLpp3p proteins, S. cerevisiae Dpp1p and Lpp1p proteins, all available mammalian lipid phosphate phosphatase proteins, Drosophila Wunen protein, as well as uncharacterized putative lipid phosphate phosphatase-like proteins identified in A. thaliana, D. melanogaster, C. elegans, and Schizosaccharomyces pombe genomes, were used for comparison. The tree was built using the neighbor joining method, wherein point accepted mutation (PAM) distances were computed based on reliably aligned sites. The SwissProt accession numbers (in brackets) designate all protein sequences. The length of horizontal branches is such that the evolutionary distance between two proteins is proportional to the total length of the horizontal branches that connect them. Bootstrap values are shown at the nodes. H.s. Lpp1p, Lpp2p, and Lpp3p: Homo sapiens Lpp1p, Lpp2p, and Lpp3p; S.c.Dpp1p: Saccharomyce scerevisiae Dpp1p; S.p.Dpp1p: Schizosaccharomyces pombe Dpp1p; R.n. Lpp1p and Lpp3p: Rattus norvegicus Lpp1p and Lpp3p; M.m. Lpp1p and Lpp2p: Mus musculus Lpp1p and Lpp2p; D.m.: D. melanogaster lipid phosphate phosphatase; C.e.: C. elegans lipid phosphate phosphatase; C.p.Lpp1p: Cavia porcellus Lpp1p

3.2. mRNA level of CrPAP2 under N-sufficient and N-limited conditions

To determine the mRNA levels of CrPEPC1-1, we collected cells of Chlamydomonas (2×10⁶ cells/ml) by centrifugation. After washing, the cells were transferred to the HSM or HSM-N medium for further cultivation. The algal cells that were harvested at 24, 48, 72, or 96 h were used for RNA extraction. We quantitatively determined the expression of CrPAP2 gene in these samples using reverse transcription and real-time PCR. Results indicate that the cells accumulated 2–5 times more lipid under the −N condition than the +N condition (Fig. 2). In addition, the mRNA levels of CrPAP2 increased significantly. Thus, we investigated whether the increase in the level of mRNA of CrPAP2 influenced lipid accumulation.

Fig. 2.

Biomass (a), lipid content (b), and mRNA levels (c) of CrPAP2 in HSM and HSM-N medium

The mRNA levels of C. reinhardtii CC425 samples grown in the indicated medium for 1, 2, 3, or 4 d, were analyzed using RT-PCR. +N: cells cultivated in N-sufficient HSM medium; −N: cells cultivated in N-free HSM medium. Data are expressed as mean±SD (n=3)

3.3. Silencing of CrPAP2 decreased lipid content in C. reinhardtii

To determine further the relationship between CrPAP2 expression and lipid accumulation, we examined the effects of artificial silencing of the CrPAP2 gene on the lipid content of C. reinhardtii. Based on the CrPAP2 (343983) sequences retrieved from the JGI C. reinhardtii v4.0 database, we designed primers to amplify the fragment of the coding region of CrPAP2. The DNA fragment was subcloned and then used to generate the CrPAP2 RNAi construct pMaa7IR/CrPAP2 IR. After transforming the silencing construct into C. reinhardtii CC425, more than 150 positive transformants were obtained. We selected three transgenic algae for measuring the lipid content and mRNA levels of the target gene. Strains transformed with the vector pMaa7IR/XIR were used as controls. Analysis of the transgenic algae revealed that the lipid content decreased by 12.4%–17.4% after 6 d of cultivation (Fig. 3b), although no differences in their growth were found (Fig. 3a). To evaluate the effectiveness of our RNAi constructs, we analyzed the abundance of target gene-specific mRNA using real-time PCR in transgenic algae. The level of the mRNA of CrPAP2 decreased to 83.4% from 94.0% (Fig. 3c), indicating high-efficiency silencing by these constructs.

Fig. 3.

Biomass (a), lipid content (b), and mRNA levels (c) of CrPAP2 RNAi transgenic algae strains

Maa7-4 (10, 19), pMaa7IR/XIR transgenic algae strains; PAP2-RNAi-3 (10, 56), pMaa7IR/CrPAP2 IR transgenic algae strains. Data are expressed as mean±SD (n=3)

Similar results were obtained from Nile Red staining. Based on microscopic analysis, fewer oil droplets with yellow fluorescence were found in CrPAP2 RNAi transgenic algae than in the pMaa7IR/XIR transgenic algae (Fig. 4). These data indicate that the regulation of CrPAP2 gene expression can affect the cell lipid content.

Fig. 4.

Microscopic observation of CrPAP2 transgenic algae strains (250× Nikio 80i)

Above: bright field, for cell morphology; Below: dark field, for oil droplets. After 4 d of cultivation in HSM medium, fewer oil droplets of CrPAP2 RNAi transgenic algae were found. Maa7-19, pMaa7IR/XIR transgenic algae strain number 19; PAP2-RNAi-56, pMaa7IR/CrPAP2 IR transgenic algae strain number 56

3.4. Over-expression of CrPAP2 improved lipid content in C. reinhardtii

The decrease in lipid content caused by RNAi silencing of CrPAP2 suggested that the expression of the CrPAP2 gene affected the biosynthesis of triglycerides in C. reinhardtii. Thus, we examined whether CrPAP2 over-expression could increase the lipid content in C. reinhardtii. The pCAMPAP2 vectors that expressed the CrPAP2 gene from the CAMV 35S promoter were introduced into C. reinhardtii. The lipid contents and growth rates of three randomly selected transgenic algae were determined in each transgenic algae line. Over-expression of the CrPAP2 gene increased growth rate in the early stages from Days 2 to 5 (Fig. 5a). Compared with the control pCAMBIA1302 transgenic algae lines, the over-expression of CrPAP2 increased lipid content. For instance, after 6 d of growth in HSM medium, the lipid contents of transgenic algae increased by 7.5%–21.8% (Fig. 5b). Compared with pCAMBIA1302 transgenic stains, the mRNA levels of CrPAP2 increased by 329%–523% (Fig. 5c). Moreover, analysis of the PAP activities of the transgenic lines showed a modest increase in PAP activity (Fig. 5d). The increase in lipid content was also observed using Nile Red dye staining (Fig. 6). More oil droplets were found in CrPAP2 over-expression transgenic algae compared with the controls, as determined by microscopic analysis. These data indicate that an increase in CrPAP2 gene expression improved cell lipid content.

Fig. 5.

Biomass (a), lipid content (b), mRNA level (c), and PAP activity (d) assays of CrPAP2 over-expression transgenic algae in HSM medium

pCAMBIA-2 (8, 16), pCAMBIA1302 transgenic algae strains; pCAMBIA-PAP2-4 (26, 60), pCAMPAP2 transgenic algae strains. Data are expressed as mean±SD (n=3)

Fig. 6.

Lipid content in a transgenic algae line detected by Nile Red staining (250× Nikio 80i)

Above: bright field, for cell morphology; Below: dark field, for oil droplets. After 4 d of cultivation in HSM medium, more oil droplets of CrPAP2 transgenic algae were found. pCAMBIA-16, pMCAMBIA1302 transgenic algae strain number 16; pCAMBIA-PAP2-60, pCAMPAP2 transgenic algae strain number 60

3.5. Expression of CrPAP2 in E. coli BL21 and detection of in vitro enzyme activity

The plasmid pGEX-6p-1-PAP2 was constructed to express CrPAP2 genes in E. coli to verify their enzyme activities. The recombinant vector was transformed into the E. coli BL21 strain. Transformants were grown in lysogeny broth medium and induced with 1 mmol/L of isopropylthiogalactoside. The supernatant was then loaded onto a 15% SDS-PAGE gel. A GST-CrPAP2 protein band of 60 kD was observed (Fig. 7a). The fusion proteins were purified using columns, followed by enzyme activity assay (Figs. 7b and 7c). Compared with the control, the enzyme activity of CrPAP increased by 27- to 29-fold (Fig. 7c). This behavior indicated that the cloned CrPAP2 gene exhibits biological activities.

Fig. 7.

Expression of CrPAP2 in E. coli BL21 and in vitro enzyme activity assay

After induction by IPTG and cultivation for 0, 2, or 4 h, total protein was harvested and run on SDS-PAGE. (a) Expression of CrPAP2 in E. coli BL21; (b) The purified GST-CrPAP2; (c) In vitro enzyme activity assay of GST-CrPAP2. GST, E. coli BL21 transformed with pGEX-6p-1; GST-CrPAP2, E. coli BL21 transformed with plasmid pGEX-6p-1-PAP2 to express the fusion protein GST-CrPAP2. The induction time of the cultured cells is indicated by 0, 2, 4, and 6 h. The GST and GST-CrPAP2 fusion proteins are indicated by the arrow

4. Discussion

PAP is a very important enzyme in lipid biosynthesis. This enzyme cleaves the phosphomonoester bond present in PA, yielding DAG and Pi (Smith et al., 1957; Carman, 1997). PAP acts as a pivotal biocatalyst in the metabolic flux between the different classes of glycerolipids within the endoplasmic reticulum and has an important role in the phospholipase D signaling pathway (Exton, 1994). Several PAP enzymes have been described from S. cerevisiae, mammalian, and Arabidopsis cells (Phan and Reue, 2005; Hans et al., 2006). These enzymes can utilize a variety of lipid phosphate substrates in vitro, including lysophosphatidic acid, phosphatidic acid, diacylglycerol pyrophosphate, sphingosine 1-phosphate, and ceramide 1-phosphate, to form DAG. They are important in lipid accumulation and cell signal transductions. In the Kennedy pathway of TAG synthesis, formation of DAG is the integral step. Moreover, DAG is not only essential for TAG formation, but is also a substrate in the synthesis of phosphatidylethanolamine (PE) and phosphatidylcholine (PC). In Chlamydomonas, instead of PC, the betaine lipid diacylglyceryl-N,N,N-trimethylhomoserine (DGTS) is the predominant glycerolipid building block of membranes (Klug and Benning, 2001). Therefore, the up- or down-regulation of PAP activities affects the content of DAG, which also affects the content of TAG, PE, and DGTS. In this study, we identified a new Chlamydomonas phosphate phosphatase gene, CrPAP2, which is different from genes in the mammalian or Arabidopsis phosphate phosphatase gene families. The in vitro expression protein has the same function as phosphate phosphatase, namely, it catalyzes the reaction of phosphatidic acid to yield diacylglycerol and Pi. We showed that an increase or decline in the expression of the gene could affect the lipid content of algal cells. Our findings suggest that increasing oil content can result from the over-expression of CrPAP2 in microalgae. Further research on CDP-ethanolamine:diacylglycerol ethanolamine phosphotransferases and betaine lipid synthase will help in understanding TAG metabolism and phospholipid biosynthesis in photosynthetic eukaryotic cells.

List of electronic supplementary materials

Fig. S1

Correlation between biomass (cell dry weight g/L) and the optical density OD₄₉₀

Fig. S2

Correlation between lipid concentration (triolein mg/20 ml) and the fluorescence value FD_470/570