Generation of N-Acylphosphatidylethanolamine by Members of the Phospholipase A/Acyltransferase (PLA/AT) Family*

Background: The mammalian enzymes that form N-acylphosphatidylethanolamines (NAPEs), precursors of bioactive N-acylethanolamines, are poorly understood. Results: PLA/AT family proteins, previously known as tumor suppressors, catalyzed N-acylation of phosphatidylethanolamine, and their overexpression in animal cells remarkably increased endogenous levels of NAPEs. Conclusion: These proteins may function as NAPE-forming enzymes in vivo. Significance: Our results may contribute to a better understanding of the regulatory mechanisms of N-acylethanolamine levels. Bioactive N-acylethanolamines (NAEs), including N-palmitoylethanolamine, N-oleoylethanolamine, and N-arachidonoylethanolamine (anandamide), are formed from membrane glycerophospholipids in animal tissues. The pathway is initiated by N-acylation of phosphatidylethanolamine to form N-acylphosphatidylethanolamine (NAPE). Despite the physiological importance of this reaction, the enzyme responsible, N-acyltransferase, remains molecularly uncharacterized. We recently demonstrated that all five members of the HRAS-like suppressor tumor family are phospholipid-metabolizing enzymes with N-acyltransferase activity and are renamed HRASLS1–5 as phospholipase A/acyltransferase (PLA/AT)-1–5. However, it was poorly understood whether these proteins were involved in the formation of NAPE in living cells. In the present studies, we first show that COS-7 cells transiently expressing recombinant PLA/AT-1, -2, -4, or -5, and HEK293 cells stably expressing PLA/AT-2 generated significant amounts of [14C]NAPE and [14C]NAE when cells were metabolically labeled with [14C]ethanolamine. Second, as analyzed by liquid chromatography-tandem mass spectrometry, the stable expression of PLA/AT-2 in cells remarkably increased endogenous levels of NAPEs and NAEs with various N-acyl species. Third, when NAPE-hydrolyzing phospholipase D was additionally expressed in PLA/AT-2-expressing cells, accumulating NAPE was efficiently converted to NAE. We also found that PLA/AT-2 was partly responsible for NAPE formation in HeLa cells that endogenously express PLA/AT-2. These results suggest that PLA/AT family proteins may produce NAPEs serving as precursors of bioactive NAEs in vivo.

Ethanolamides of different long-chain fatty acids constitute a class of naturally occurring lipid molecules and are collectively referred to as N-acylethanolamines (NAEs) 2 (1,2). NAEs show a wide variety of biological activities depending on their acyl chains, and these activities are based on their abilities to bind to and activate specific receptors. In particular, N-arachidonoylethanolamine, also known as anandamide, has attracted much attention as an endogenous ligand for cannabinoid receptors (3) and for transient receptor potential vanilloid type-1 (TRPV1) (4). N-Palmitoylethanolamine and N-oleoylethanolamine have been documented as an anti-inflammatory and analgesic substance (5,6) and appetite-suppressing substance (7), respectively, through peroxisome proliferator-activated receptor (PPAR) ␣ (8,9). N-Oleoylethanolamine was also reported to be an agonist of TRPV1 (10) and GPR119 (11).
NAE is biosynthesized from membrane glycerophospholipids by two steps of enzyme reactions ( Fig. 1) (12,13). The first step of the NAE-biosynthesizing pathway is the formation of N-acylphosphatidylethanolamine (NAPE) by transferring an acyl chain from the sn-1 position of glycerophospholipid to the amino group of phosphatidylethanolamine (PE). Although it has been established that membrane-bound Ca 2ϩ -dependent N-acyltransferase (Ca-NAT) catalyzes this reaction in the brain (14 -16), the enzyme has not been purified or cloned. The sec-ond step is the hydrolysis of NAPE to NAE by NAPE-hydrolyzing phospholipase D (NAPE-PLD) (17). Alternative pathways independent of NAPE-PLD are also known (18 -20). As a major degradative pathway of NAE, the hydrolysis of NAE to fatty acid and ethanolamine is catalyzed by membrane-bound fatty acid amide hydrolase (FAAH) or lysosomal NAE-hydrolyzing acid amidase (NAAA) (Fig. 1) (12,21,22). Recent studies using knock-out mice and specific inhibitors revealed that these NAE-hydrolyzing enzymes are promising targets for the development of therapeutic drugs (6,23).
The HRAS-like suppressor family (also known as the H-rev107 family) consists of tumor suppressor genes negatively regulating the activity of oncogene Ras (16, 24 -27). In human beings, five members (HRASLS1-5) belong to this family. Recently, we demonstrated that the gene products of all five members possess phospholipase A 1/2 (PLA 1/2 ) activity, which releases fatty acid from the sn-1 or sn-2 position of glycerophospholipid, and O-acyltransferase activity, which transfers an acyl group from glycerophospholipid to the hydroxyl group of lysophospholipid (28 -31). Based on these phospholipid-metabolizing activities, we proposed to designate the gene products of HRASLS1-5 as phospholipase A/acyltransferase-1-5 (PLA/ AT-1-5), respectively (31). We will use these new names throughout this report.
Expression of PLA/AT Family Members in Animal Cells-COS-7 cells were grown at 37°C to 90% confluency in 100-mm dishes containing Dulbecco's modified Eagle's medium with 10% fetal bovine serum in a humidified 5% CO 2 and 95% air incubator. The expression vectors harboring N-terminally FLAG-tagged PLA/AT-1-5 or the insert-free pEF1/Myc-His vector were introduced into COS-7 cells using Lipofectamine 2000 according to the manufacturer's instructions. Forty eight hours after transfection, cells were harvested, sonicated three times each for 3 s in 20 mM Tris-HCl (pH 7.4), and used for enzyme assays. For the experiments shown in Fig. 4, recombinant FLAG-tagged PLA/AT-2 was purified by anti-FLAG M2 affinity chromatography as described previously (30). For the stable expression of PLA/AT-2, HEK293 cells were transfected with pEF1/Myc-His vector harboring N-terminally FLAGtagged PLA/AT-2 or the insert-free pEF1/Myc-His vector using Lipofectamine 2000. Cells were selected in the medium containing 1 mg/ml geneticin. Clonal cell lines PLA/AT-2-H and PLA/AT-2-L were isolated by colony lifting, and propagated. PLA/AT-3-expressing HEK293 cells were established previously (36).
RNA Interference-siRNAs were introduced into PLA/AT-2-H cells or HeLa cells with Lipofectamine RNAiMAX according to the manufacturer's instructions. The final concentration of siRNA was 20 nM. Forty eight hours after transfection, cells were subjected to RT-PCR, the N-acyltransferase assay, metabolic labeling with [ 14 C]ethanolamine, or LC-MS/MS analysis.
Metabolic Labeling-Cells were grown at 37°C to 80% confluency in a 100-mm dish containing Dulbecco's modified Eagle's medium with 10% fetal calf serum and were labeled with [ 14 C]ethanolamine (1.6 Ci) or [ 14 C]palmitic acid (1.6 Ci) for 18 h. Cells were then harvested and washed twice with PBS. Total lipids were extracted by the method of Bligh and Dyer (34), spotted on a silica gel thin layer plate (20-cm height), and developed at 4°C for 90 min in solvent A. The distribution of radioactivity on the plate was visualized and quantified using a BAS1500 bioimaging analyzer.
Western Blotting-Cells were homogenized in homogenization buffer (0.25 M sucrose, 1 mM EDTA, and 20 mM HEPES (pH 7.4)) by being passed through a 27-gauge syringe (37,38), and nuclei and unbroken cells were removed by centrifugation at 800 ϫ g for 10 min at 4°C. Postnuclear supernatant fractions were then centrifuged at 105,000 ϫ g for 30 min at 4°C to separate the cytosol (supernatant fractions) from cellular organelles (particulate fractions). Samples were separated by SDS-PAGE and electrotransferred to a hydrophobic polyvinylidene difluoride membrane (Hybond P). The membrane was blocked with PBS containing 5% dried milk and 0.1% Tween 20 (buffer A) and then incubated with primary antibodies (1:2000 dilution) in buffer A at room temperature for 1 h, followed by incubation with horseradish peroxidase-labeled secondary antibodies (1:4000 dilution) in buffer A at room temperature for 1 h. Proteins were finally treated with an ECL Plus kit and visualized with the aid of a LAS1000plus lumino-imaging analyzer (FUJIX Ltd.).
Lipid Analysis by LC-MS/MS-Lipids were extracted from cells by a modification of the method of Bligh and Dyer, essentially as described previously (39). In this method, cells were suspended in 3.8 ml of a mixture of chloroform, methanol, 0.07 M KCl (1:2:0.8, v/v) on ice followed by sonication for 10 -20 s. A mixture of internal standards for LC-MS/MS was added to this suspension. After standing for 20 min on ice, the mixture was centrifuged at 1400 ϫ g for 10 min. The supernatant was withdrawn, and the resultant pellet was mixed with 1.9 ml of chloroform/methanol/water (1:2:0.8) followed by centrifugation. Supernatants were combined, and 1.5 ml each of chloroform and water was added to the sample to produce phase separation. After centrifugation of the mixture, the organic lower phase was withdrawn. The upper layer was mixed with 3 ml of chloroform/methanol (17:3), and the mixture was centrifuged. Combined lower layers were evaporated to dryness under a stream of nitrogen gas, and half of this lipid extract was recon-stituted in 0.1 ml of methanol/water (95:5, v/v) containing 0.05 M ammonium formate in an insert of a brown glass vial for LC-MS/MS. The lipid/phosphate concentration in another half of the lipid extract was determined as described previously (40).
LC-MS/MS was performed on a quadrupole-linear ion trap hybrid MS, 4000 Q TRAP (Applied Biosystems/MDS Sciex, Concord, Ontario, Canada) with a 1100 LC system (Agilent Technologies, Wilmington, DE) combined with an HTS-PAL autosampler (CTC Analytics AG, Zwingen, Switzerland), essentially as described previously (20). The extract was analyzed for molecular species of NAPE by LC on an Imtakt Unison UK-Amino column (100 ϫ 2 mm, 3.0-m particle size) at a flow rate of 0.1 ml/min. The mobile phase was a mixture of acetonitrile/methanol (95:5, v/v) containing 0.1% triethylamine. The molecular species of glycerophospho-N-acylethanolamine (GP-NAE) and NAE in the extract were separated on a Supelco Ascentis Express C18 reverse phase column (100 ϫ 2.1 mm, 2.7-m particle size) with methanol/water (95:5) containing 5 mM ammonium formate at a flow rate of 0.20 (for GP-NAE) or 0.15 ml/min (for NAE). The molecular species of PE were separated by LC on Cadenza CD-C18 (100 ϫ 1.0 mm, 3-m particle) with acetonitrile/methanol (1:1) containing 5 mM ammonium formate at a flow rate of 0.15 ml/min. Routinely, 5-l aliquots of the test solution in an insert were applied using the autosampler. In the negative ion mode of operation with multiple reaction monitoring, [R 2 COO] Ϫ and the deprotonated molecular ion for NAPE were selected for Q3 and Q1. In the positive ion mode of operation with multiple reaction monitoring, [ethanolamine] ϩ at m/z 62 for NAE and [M ϩ H Ϫ phosphoethanolamine] ϩ for PE were selected as Q3 in combination with the protonated molecular ion as Q1. The molecular species of NAE, NAPE, and diacyl-PE were quantified using deuterated N-palmitoylethanolamine, N-heptadecanoyl-1,2-dipalmitoyl-PE, and 1,2-dimyristoyl-PE as internal standards, respectively. Corrections were made for the quantification of molecular species of N-acylated plasmenylethanolamine (pNAPE) and plasmenylethanolamine based on the slopes of the calibration lines constructed with N-heptadecanoyl- respectively. Negative ions due to the molecular species of GP-NAE were quantified at a combination of deprotonated molecular ions and [glycerophosphate (171)] Ϫ for Q1 and Q3, based on the peak ratios relative to glycerophospho-N-heptadecanoylethanolamine. Values are represented as picomoles/ mol or nanomoles/mol of total phospholipids.

Generation of NAPE in Cells Overexpressing PLA/AT Family
Members-To examine whether PLA/AT family members can generate NAPE in living cells, we transiently expressed each of the PLA/AT-1-5 in COS-7 cells. Expression was confirmed by Western blotting with an anti-FLAG antibody, which recognized the FLAG tag attached to recombinant PLA/AT-1-5 (data not shown). Homogenates of the transfectants were allowed to react with 1,2-[ 14 C]dipalmitoyl-PC and dioleoyl-PE as an acyl donor and an acyl acceptor in the N-acyltransferase reaction, respectively, and products were separated by TLC. The results showed the formation of a radioactive band corresponding to N-palmitoyl-PE by all of PLA/AT-1-5 ( Fig. 2A). A radioactive band corresponding to free palmitic acid was also detected in each PLA/AT as the PLA 1/2 reaction product. The ratio of N-acyltransferase activity to PLA 1/2 activity was largely different among PLA/AT-1-5 (Fig. 2B). The highest ratio (5.6) was seen with PLA/AT-2, followed by PLA/AT-5 and -1. Conversely, PLA/AT-3 and -4 showed lower ratios (Ͻ1). A radioactive band comigrated with authentic N-palmitoyl-lyso-PE also being detected. On the other hand, N-palmitoyl-PE and N-palmitoyl-lyso PE were hardly detectable with the homogenate of COS-7 cells transfected with the insert-free vector.
Next, these cells were metabolically radiolabeled with [ 14 C]ethanolamine, and total lipids extracted from cells were analyzed by TLC. As shown in Fig. 2C, radioactive bands that comigrated with authentic N-palmitoyl-PE, N-palmitoylethanolamine, and N-palmitoyl-lyso-PE were clearly detected in most PLA/AT-expressing cells. The radioactive substance corresponding to NAPE was extracted from the silica gel plate and treated with recombinant NAPE-PLD, an enzyme that specifically hydrolyzes NAPE to NAE and phosphatidic acid. This treatment led to the production of a radioactive band that comigrated with authentic N-palmitoylethanolamine (Fig. 2F), confirming that the original band is NAPE. When radioactive NAPE was quantified (Fig. 2D), high levels of NAPE (19.4 and 8.9% of total radioactivity) were detected in PLA/AT-2-expressing cells and PLA/AT-1-expressing cells, respectively, followed by PLA/AT-4-expressing cells and PLA/AT-5-expressing cells. Although PLA/AT-3 cells generated a small amount of NAPE, its level was not significantly different from that in control COS-7 cells. The content of NAE also showed a similar tendency with the highest level (1.6% of total radioactivity) in PLA/AT-2 cells (Fig. 2E). We also labeled these cells with [ 14 C]palmitic acid and confirmed the generation of [ 14 C]NAPE in cells expressing PLA/AT-1, -2, -4, or -5, with the highest NAPE radioactivity in PLA/AT-2 cells (Fig. 2G). These results showed that PLA/AT-1, -2, -4, and -5 have the capability to generate NAPE in living cells. Because PLA/AT-2-expressing cells showed the highest levels of NAPE and NAE, we focused on the characterization of PLA/AT-2 hereafter.
NAPE Formation by PLA/AT-2 Requires Its Enzyme Activity-Because Cys-113 of PLA/AT-2 is presumed to be a catalytic nucleophile (30), its C113S mutant was expected to be catalytically inactive. We thus constructed this mutant, transiently expressed it in COS-7 cells, and confirmed its expression by Western blotting (Fig. 3A). The cell homogenate was essentially free of N-acyltransferase (Fig. 3B), and metabolic labeling of C113S-expressing cells with [ 14 C]ethanolamine did not increase levels of radioactive NAPE (Fig. 3C) and NAE (Fig. 3D). These results strongly suggest that the enzyme activity of PLA/ AT-2 is required for the production of NAPE and NAE in PLA/ AT-2-expressing cells.
PLA/AT-2 Preferentially Transfers sn-1 Acyl Chain of Phospholipid to PE-We earlier found that the purified recombinant PLA/AT-2, which functions as a PLA 1/2 enzyme, preferentially Formation of N-Acylphosphatidylethanolamine by PLA/AT Family SEPTEMBER 14, 2012 • VOLUME 287 • NUMBER 38 releases sn-1 fatty acid over sn-2 fatty acid from glycerophospholipids (30). To determine which acyl chain is utilized in the N-acyltransferase reaction, we allowed the purified PLA/AT-2 to react with 1-  (Fig. 5A). mRNA of PLA/AT-2 was negligible in HEK293 cells transfected with the insert-free vector. N-Acyltransferase activity in the homogenate of PLA/AT-2-H cells was about 2-fold higher than that of PLA/AT-2-L cells (Fig. 5, B and C). When 1 mM EDTA was replaced with 1 mM Ca 2ϩ , activity did not increase but decreased by about 50% (Fig. 5C). When the homogenate of PLA/AT-2-H cells was centrifuged at 105,000 ϫ g, both soluble and particulate fractions showed N-acyltransferase activity with a 1.4-fold higher activity in the soluble fraction (Fig. 5D).
To examine the intracellular generation of NAPE, PLA/AT-2-L and PLA/AT-2-H cells were metabolically radiolabeled with [ 14 C]ethanolamine. When total lipids were extracted from cells and separated by TLC, NAPE levels in these cells were 7.7and 14.1-fold higher than that of control HEK293 cells, respectively (Fig. 6, A and B). NAE levels also increased 2.8-and 4.6-fold (Fig. 6B). Thus, cellular levels of NAPE and NAE in these two PLA/AT-2-expressing cells correlated well with PLA/AT-2 mRNA levels and N-acyltransferase activities in the homogenates.
To rule out the possibility that the accumulation of NAPE was caused by the insertion of the PLA/AT-2 gene into a specific region of the genome that contains one or more genes related to the metabolism of NAPE, we suppressed the expression of recombinant PLA/AT-2 in PLA/AT-2-H cells by two    siRNAs targeting different regions of PLA/AT-2. As shown in Fig. 7A, RT-PCR analysis revealed a strong suppression of PLA/ AT-2 mRNA expression in cells treated with PLA/AT-2 siRNAs. N-Acyltransferase activities in the homogenates of PLA/AT-2 knockdown cells were reduced to 11-19% that of cells treated with a control siRNA (Fig. 7B). When knockdown cells were labeled with [ 14 C]ethanolamine, NAPE levels almost completely reverted to that of control cells (Fig. 7C). These results confirmed that the stable expression of PLA/AT-2 causes the accumulation of NAPE in living HEK293 cells by its N-acyltransferase activity.
Analysis of NAPEs, pNAPEs, and Their Metabolites by LC-MS/MS-We next analyzed the molecular species of N-acylated ethanolamine phospholipids and their metabolites in PLA/AT-2-H cells and control HEK293 cells by LC-MS/MS. Concerning N-acylated ethanolamine phospholipids, we measured both the diacyl-type (NAPE) and plasmalogen-type (pNAPE) (Fig. 8). We confirmed the preferential liberation of the sn-2 fatty acyl group over the sn-1 fatty acyl group from a standard N-heptadecanoyl-PE (1-palmitoyl-2-oleoyl) under our conditions for tandem mass spectrometry. However, we could not exclude a possibility that [R 1 COO] Ϫ was partially involved in the ion peaks assigned as [R 2 COO] Ϫ when endogenous NAPE species were analyzed. Thus, we tentatively assigned the molecular species of NAPE (diacyl) in terms of both the combined chain length and unsaturation degree of the sn-1 O-and N-linked fatty acyl moieties together with those of sn-2 O-linked fatty acyl moiety. As shown in Fig. 8A, the expression of PLA/AT-2 remarkably increased endogenous levels of most species of NAPE. Major sn-1 O-acyl ϩ N-acyl species of NAPE in PLA/AT-2-H cells were 32:0, 32:1, 34:0, 34:1, 36:0, 36:1, and 36:2. The total amount of NAPE species in cells was 13-fold larger than that in control cells. The sn-2 O-acyl chains of NAPE were mostly 18:1. However, the amounts of most pNAPE species were slightly increased or almost unaltered by the expression of PLA/AT-2 (Fig. 8B). The total amount of pNAPE species in PLA/AT-2-H cells was only 1.4-fold larger than that in control cells.
As shown in Fig. 9A, the major NAEs were 18:0, 16:0, and 18:1. The total amount of NAEs was 12-fold higher in PLA/ AT-2 cells. A small amount of anandamide was detected only in PLA/AT-2-H cells. GP-NAE is an intermediate in the NAPE-PLD-independent pathway, which forms NAE from NAPE (20,41). Various species of GP-NAE were remarkably increased by the overexpression of PLA/AT-2 (Fig. 9B). Major N-acyl species of GP-NAE were also 18:0, 16:0, and 18:1. These results showed that the expression of PLA/AT-2 causes remarkable increases in not only NAPEs but also their metabolites NAEs and GP-NAEs.
Peroxisomal Dysfunction of PLA/AT-2-H Cells-Our recent study revealed that the overexpression of PLA/AT-3 (H-rev107) results in a drastic decrease in the levels of ethertype lipids, including plasmenylethanolamine (36). We also found the dysfunction of peroxisomes, organelles involved in the biosynthesis of ether-type lipids, in PLA/AT-3-expressing cells. We therefore examined whether overexpression of PLA/ AT-2 also decreases endogenous plasmenylethanolamine levels. As analyzed by LC-MS/MS (Fig. 10B), all species of plasmenylethanolamine were reduced in PLA/AT-2-H cells. The total amount of plasmenylethanolamine species was decreased by 91%. However, all species of diacyl-type PE were increased by the overexpression of PLA/AT-2 (Fig. 10A). Reductions in plasmenylethanolamine as a precursor of pNAPE may explain why pNAPE species were almost unaltered or slightly increased by the expression of PLA/AT-2 in contrast to the marked increase in NAPE species.
We also examined the expression and subcellular localization of PMP70 and catalase, two representative peroxisomal proteins, by Western blotting (Fig. 10C). In control cells, these proteins were localized in the particulate fraction as expected. However, in PLA/AT-2-H cells, PMP70 was hardly detected in the particulate or supernatant fraction, and catalase was detected mostly in the supernatant fraction. In contrast to their abnormal localization, RT-PCR analysis confirmed normal expression levels of PMP70 and catalase mRNAs in PLA/AT-  (Fig. 10D). Similar results were obtained with the peroxisomal proteins in PLA/AT-3-expressing cells (Fig. 10, C and  D), in agreement with our previous report (36). These results strongly suggested that the expression of PLA/AT-2 as well as PLA/AT-3 causes the dysfunction of peroxisomes.

2-H cells
Expression of Related Enzymes in PLA/AT-2-expressing Cells-NAPE-PLD is a major enzyme responsible for the generation of NAE from NAPE. Overexpression of NAPE-PLD in mammalian cells leads to an increase in NAE levels with a concomitant decrease in NAPE levels (35). To examine whether the NAPE generated in PLA/AT-2-H cells can be metabo-lized by NAPE-PLD, we transiently expressed NAPE-PLD in PLA/AT-2-H cells. The homogenate of cells showed a NAPE-PLD activity as high as 27.1 nmol/min/mg protein, which produced N-palmitoylethanolamine from N-palmitoyl-PE (Fig. 11, A and B). Metabolic labeling with [ 14 C]ethanolamine revealed that the expression of NAPE-PLD results in a strong reduction in NAPE levels (Fig. 11, C and D). The concomitant increase in NAE levels was observed only in the presence of URB597, an FAAH inhibitor (Fig. 11, C and D) (42), suggesting rapid degradation of NAE by endogenous FAAH.

Formation of N-Acylphosphatidylethanolamine by PLA/AT Family
Next, we simultaneously expressed both NAPE-PLD and NAAA, an NAE-hydrolyzing enzyme different from FAAH (12), in PLA/AT-2-H cells. The transient expression of NAAA brought a high N-palmitoylethanolamine-hydrolyzing activity in the cell homogenate at pH 4.5, which is optimal for NAAA, whereas that of control cells expressing only NAPE-PLD was almost inactive under the same conditions (Fig. 12A). When cells were radiolabeled with [ 14 C]ethanolamine in the presence of URB597, NAE levels in NAAA-expressing cells decreased by 56% relative to control cells (Fig. 12, B and C). These results indicated that NAPE generated by PLA/AT-2 can be metabolized by NAPE-PLD and that the resultant NAE is degraded by FAAH and NAAA.
PLA/AT-2-dependent Formation of NAPE Is Not Enhanced by Cellular Stimuli-It was previously reported that Ca 2ϩ ionophores A23187 and ionomycin increased NAPE levels in cortical neurons of rats and mice (14,43). This Ca 2ϩ -dependent formation of NAPE was attributed to Ca-NAT. Although NAPE formation by PLA/AT-2 was Ca 2ϩ -independent (Fig.  5C), we were curious as to whether NAPE formation by PLA/ AT-2 in living cells was stimulated by Ca 2ϩ ionophores or other reagents. PLA/AT-2-H cells were labeled with [ 14 C]ethanol-amine and then treated with A23187, forskolin (an adenylyl cyclase activator), or PMA (a protein kinase C activator) (Fig.  13). However, none of these reagents caused a significant increase in the levels of NAPE and NAE.
Possible Involvement of Endogenous PLA/AT-2 in NAPE Formation in HeLa Cells-HeLa cells endogenously express PLA/ AT-2 (30). Therefore, we were interested in whether or not endogenous PLA/AT-2 was involved in NAPE formation in HeLa cells. We confirmed the expression of PLA/AT-2 mRNA by RT-PCR (Fig. 14A). mRNAs of PLA/AT-3 and -4 were also detected (Fig. 14A), whereas PLA/AT-1 and -5 were not detectable (data not shown). Introduction of two different siRNA constructs against PLA/AT-2 expectedly caused a decrease in PLA/AT-2 mRNA levels without affecting PLA/AT-3 and -4 levels (Fig. 14A). Both of the siRNA constructs reduced cellular levels of several species of NAPEs (Fig. 14B) and pNAPEs (Fig.  14C) in a similar manner. These results suggest that endogenous PLA/AT-2 is partly responsible for NAPE and pNAPE formation in HeLa cells.

DISCUSSION
NAPE is a class of endogenous glycerophospholipids and is well known to be precursors of bioactive NAEs (1, 2). The major route for NAPE formation in animal tissues is N-acylation of PE using glycerophospholipid as an acyl donor (13). The responsible enzyme Ca-NAT, however, remains molecularly uncharacterized. In contrast, we found an enzyme catalyzing the same reaction in a Ca 2ϩ -independent manner, and we termed it Ca 2ϩ -independent N-acyltransferase (iNAT, referred to as PLA/AT-5 in this study) (16,28). Furthermore, we reported that other members of the PLA/AT family possess PE N-acyltransferase activity together with PLA 1/2 and lysophospholipid O-acyltransferase activities (30,31). In humans, PLA/AT family members include A-C1 (PLA/AT-1), HRASLS2 (PLA/AT-2), H-rev107 (PLA/AT-3), tazarotene-induced protein 3 (TIG3, PLA/AT-4), and iNAT (PLA/AT-5). However, their roles in the in vivo formation of NAPE were poorly understood.
In this study, we first showed that transient expressions of PLA/AT-1, -2, -4, and -5 in COS-7 cells caused intracellular accumulation of NAPE. PLA/AT-3-expressing cells did not show a significant increase in NAPE levels, although the cell homogenate showed N-acyltransferase activity. The highest NAPE level in PLA/AT-2-expressing cells was consistent with the highest N-acyltransferase activity in their homogenate, and its enzymatically inactive mutant C113S failed to increase cellular NAPE levels. We have also reported that purified recombinant PLA/AT-2 showed the highest N-acyltransferase activity among purified PLA/AT family proteins (30,31). We next revealed that endogenous levels of NAPE and NAE in HEK293 cells were markedly increased by stable expression of PLA/ AT-2. With the aid of two clonal cells (PLA/AT-2-H and PLA/ AT-2-L cells), which expressed PLA/AT-2 at different levels, we showed that expression levels of PLA/AT-2 correlated well with endogenous NAPE levels in living cells as well as N-acyltransferase activity in cell homogenates. Moreover, knockdown of overexpressed PLA/AT-2 by siRNAs largely reduced both N-acyltransferase activity and endogenous NAPE levels. These results confirmed that PLA/AT-2 functions as a NAPE-forming Although all PLA/AT members function as phospholipidmetabolizing enzymes, we should note that the ratio of N-acyltransferase activity to PLA 1/2 activity is different among members (Fig. 2B). The tissue distribution of each PLA/AT is also different (28 -31). In addition, human tissues express all five members of the HRAS-like suppressor family (PLA/AT-1-5), although the genomes of rodents lack the genes of PLA/AT-2 and -4 (16, 28 -31). These findings suggest that each PLA/AT plays different roles in vivo. Considering the lack of PLA/AT-2 in rodents, PLA/AT-1 and -5 may be responsible for NAPE formation in these animals. Especially, PLA/AT-1 is abundantly expressed in the testis, skeletal muscle, heart, and brain of rats and mice (31). Thus, it is of interest to examine whether this protein is involved in NAPE formation in these tissues.
PLA/AT-2-expressing cells generated a large amount of NAPE without any cellular stimuli. Addition of A23187, forskolin, or PMA to cells did not alter intracellular NAPE levels. This was consistent with the Ca 2ϩ independency of N-acyltransferase activity in PLA/AT-2 (Fig. 5C). It is generally accepted that the formation of NAPE by N-acyltransferase is the principal rate-limiting step in the NAE-biosynthesizing pathway (13). In rat and mouse cortical neurons, Ca 2ϩ ionophores augmented the generation of NAPE (14,43), and this finding was fortified with the fact that brain N-acyltransferase (Ca-NAT) is stimulated by Ca 2ϩ (15,16). However, a certain level of NAPE appears to be present in various tissues without any cellular stimuli (2). PLA/AT-2 and other members of the PLA/AT family may thus play a role in maintaining the basal levels of NAPE. Alternatively, different from recombinant PLA/ AT-2, endogenous PLA/AT-2 may be regulated by a Ca 2ϩ -dependent protein.
The major N-acyl species of NAE in PLA/AT-2-H cells were 16:0, 18:0, and 18:1. Moreover, because the total number of double bonds in sn-1 O-acyl and N-acyl chains of NAPE was mostly 0 or 1, N-acyl species of NAPE appeared to be mostly saturated and monounsaturated acyl chains. Thus, our results using PLA/AT-2-H cells agree with the fact that NAPEs containing a saturated or monounsaturated acyl chain at the N position serve as precursors of bioactive saturated and monounsaturated NAEs such as N-palmitoylethanolamine and N-oleoylethanolamine. In contrast, anandamide (N-arachi-donoylethanolamine) and its precursor N-arachidonoyl-PE were minor components among NAEs and NAPEs, respectively. It is well known that polyunsaturated fatty acids such as arachidonic acid are mostly esterified at the sn-2 position rather than sn-1 position of glycerophospholipids. These results are in agreement with our finding that purified PLA/AT-2 transferred an acyl chain principally from the sn-1 position of PC to the amino group of PE. Thus, PLA/AT-2 does not appear to be involved in a putative anandamide-specific pathway.
PLA/AT-2-H cells showed elevated levels of NAE and GP-NAE in addition to NAPE. Because GP-NAE is an intermediate metabolite generated by double deacylation of NAPE in the NAPE-PLD-independent pathway, these results suggest that NAPE generated by PLA/AT-2 is converted to NAE directly by endogenous NAPE-PLD or through the NAPE-PLD-independent pathway via GP-NAE. Transient expression of NAPE-PLD in cells caused a remarkable decrease in NAPE levels. Concomitant increases in NAE levels were observed only in the presence of URB597, which probably inhibited endogenous FAAH. Fur-   thermore, coexpression of NAPE-PLD and NAAA (another NAE-hydrolyzing enzyme) decreased NAE levels in URB597treated PLA/AT-2-H cells. These results showed that exogenous NAPE-PLD was involved in the formation of NAE from NAPE produced by PLA/AT-2 and that exogenous NAAA as well as endogenous FAAH were utilized in the degradation of NAE in PLA/AT-2-H cells. Although we used only COS-7 and HEK293 cells to express PLA/AT-2, our results suggest that the introduction of the cDNA encoding PLA/AT-2 to various types of cells serves as a useful method to examine intracellular metabolism and the actions of NAPE and NAE as well as the intracellular effects of enzyme inhibitors.
The overexpression of PLA/AT-2 also caused a decrease in the levels of plasmenylethanolamine as well as abnormal intracellular localization of peroxisomal proteins. These results suggest that overexpressed PLA/AT-2 causes the dysfunction of peroxisomes as we recently reported in PLA/AT-3-expressing cells (36). Different from PLA/AT-2, the major catalytic activity of PLA/AT-3 is PLA 1/2 activity (29,30), and its N-acyltransferase activity is very low (Fig. 2). Therefore, it is unlikely that increased NAPE levels are responsible for peroxisomal dysfunction. We are now trying to elucidate a molecular mechanism underlying peroxisomal dysfunction in PLA/AT-2-and PLA/AT-3-expressing cells.
In conclusion, we demonstrated for the first time that PLA/ AT-2 and other PLA/AT proteins can form NAPE in living cells. We are planning further studies, including analyses of gene-disrupted and transgenic animals for PLA/AT proteins, to elucidate their physiological significance in NAPE metabolism.