Cloning and Expression of Mouse Liver Phosphatidylserine Synthase-1 cDNA
OVEREXPRESSION IN RAT HEPATOMA CELLS INHIBITS THE CDP-ETHANOLAMINE PATHWAY FOR PHOSPHATIDYLETHANOLAMINE BIOSYNTHESIS*

Scot J. StoneDagger , Zheng Cui§, and Jean E. VanceDagger

From the Lipid and Lipoprotein Research Group and Departments of Dagger  Medicine and § Biochemistry, University of Alberta, Edmonton, AB T6G 2S2, Canada

    ABSTRACT
Top
Abstract
Introduction
Procedures
Results
Discussion
References

In eukaryotic cells, phosphatidylserine (PtdSer) is synthesized by two distinct synthases on the endoplasmic reticulum by a base-exchange reaction in which the polar head-group of an existing phospholipid is replaced with serine. We report the cloning and expression of a cDNA for mouse liver PtdSer synthase-1. The deduced protein sequence is >90% identical to that of PtdSer synthase-1 from Chinese hamster ovary cells and a sequence from a human myeloblast cell line. PtdSer synthase-1 cDNA was stably expressed in M.9.1.1 cells which are mutant Chinese hamster ovary cells defective in PtdSer synthase-1 activity, are ethanolamine auxotrophs, and have a reduced content of PtdSer and phosphatidylethanolamine (PtdEtn). The growth defect of M.9.1.1 cells was eliminated, and a normal phospholipid composition was restored in the absence of exogenous ethanolamine, implying that the cloned cDNA encoded PtdSer synthase. Mouse liver PtdSer synthase-1 was also expressed in McArdle 7777 rat hepatoma cells. In addition to a 3-fold higher in vitro serine-exchange activity, these cells also exhibited enhanced choline- and ethanolamine-exchange activities and incorporated more [3H]serine into PtdSer than did control cells. However, the levels of PtdSer and PtdEtn in cells overexpressing PtdSer synthase-1 activity were not increased. Excess PtdSer produced by the transfected cells was rapidly decarboxylated to PtdEtn and the degradation of PtdSer, and/or PtdEtn derived from PtdSer, was increased. Moreover, the CDP-ethanolamine pathway for PtdEtn biosynthesis was inhibited. These data suggest that (i) cellular levels of PtdSer and PtdEtn are tightly controlled, and (ii) the metabolism of PtdSer and PtdEtn is coordinately regulated to maintain phospholipid homeostasis.

    INTRODUCTION
Top
Abstract
Introduction
Procedures
Results
Discussion
References

Phosphatidylserine (PtdSer)1 is an amino phospholipid component of all animal cell membranes, accounting for ~5-10% of membrane phospholipids. In addition to a presumed structural role in membranes, PtdSer is required for activation of protein kinase C (1) and for progression of the blood coagulation cascade (2, 3). PtdSer exposure on the cell surface also serves as a signal for recognition and removal of apoptotic cells by macrophages (4, 5). In mammalian cells, PtdSer is synthesized on ER membranes in a calcium-dependent base-exchange reaction catalyzed by PtdSer synthases (6). In this reaction, the polar head-group of an existing phospholipid, such as PtdCho or PtdEtn, is replaced by L-serine (Fig. 1). The pathway by which PtdSer is synthesized in eukaryotes is different from that in bacteria (7) and Saccharomyces cerevisiae (8, 9), in both of which PtdSer is synthesized from L-serine, with CDP-diacylglycerol as the donor of the phosphatidyl group.


View larger version (12K):
[in this window]
[in a new window]
 
Fig. 1.   Biosynthetic pathways for PtdSer and PtdEtn in mammalian cells. The abbreviations used are: PSS, PtdSer synthase; PSD, PtdSer decarboxylase; Etn, ethanolamine; DAG, diacylglycerol; EK, ethanolamine kinase; ET, CTP:phosphoethanolamine cytidylyltransferase; EPT, CDP-ethanolamine:1,2-diacylglycerol ethanolamine- phosphotransferase.

A cDNA encoding PtdSer synthase-1 was cloned from CHO-K1 cells by complementation of PSA-3 mutant cells that are defective in PtdSer synthase-1 activity (10). Expression of the cloned cDNA in PSA-3 cells showed that the enzyme catalyzed a base-exchange reaction with serine, choline, and ethanolamine. Immunoblot analysis (11) revealed that PtdSer synthase-1 is present in both microsomes and mitochondria-associated membranes (12). More recently, a cDNA that encodes PtdSer synthase-2 was also isolated from CHO-K1 cells (GenBankTM data bank accession number AB004109) (13). Although the cDNAs for PtdSer synthase-1 and -2 share little similarity, the corresponding amino acid sequences are 32% identical. Expression of PtdSer synthase-2 cDNA showed that, as predicted, this enzyme uses PtdEtn, but not PtdCho, as phosphatidyl group donor. The PtdSer synthase-2 cDNA also transformed PSA-3 cells, which lacked PtdSer synthase-1 activity, into PtdSer prototrophs (13).

M.9.1.1 cells are phenotypically similar to PSA-3 cells. In M.9.1.1 cells, the PtdSer synthase activity is ~50% that of parental CHO-K1 cells (14). Consequently, in M.9.1.1 cells that are deprived of ethanolamine, and therefore are unable to make PtdEtn via the CDP-ethanolamine pathway (Fig. 1), PtdSer and PtdEtn levels are lower than in CHO-K1 cells. The growth defect of M.9.1.1 cells is overcome when ethanolamine, PtdSer, lyso-PtdEtn, or PtdEtn are included in the culture medium. M.9.1.1 cells also do not utilize PtdCho as a substrate for PtdSer biosynthesis, although PtdEtn is converted into PtdSer almost as efficiently as in wild-type CHO-K1 cells. Therefore, the primary defect in M.9.1.1 cells was deduced to be in the PtdCho-dependent synthesis of PtdSer (14).

PtdSer synthesized on the ER is transported to mitochondria and decarboxylated to PtdEtn by the enzyme PtdSer decarboxylase (Fig. 1) (15). Nearly all mitochondrial PtdEtn in CHO cells is synthesized in situ in mitochondria by this reaction (16). PtdEtn is also synthesized on the ER by the CDP-ethanolamine pathway (Fig. 1) (17). In this pathway, ethanolamine is first phosphorylated by ethanolamine kinase to phosphoethanolamine, which is converted to CDP-ethanolamine by CTP:ethanolamine-phosphate cytidylyltransferase (ET). In the final step of the pathway, CDP-ethanolamine:1,2-diacylglycerol ethanolaminephosphotransferase transfers phosphoethanolamine from CDP-ethanolamine to diacylglycerol to produce PtdEtn. A third, and quantitatively minor, pathway for the biosynthesis of PtdEtn is the base-exchange reaction (6). The relative importance of the two major pathways for PtdEtn biosynthesis appears to depend on the cell type. In some studies, for example in rat liver (18, 19) and hamster heart (20), PtdEtn has been reported to be synthesized primarily via the CDP-ethanolamine pathway, although in another study Yeung and Kuksis (21) found that the decarboxylation of PtdSer was the major route of PtdEtn biosynthesis in rat liver. Other cells, such as CHO and BHK-21 cells, synthesize at least 80% of their PtdEtn from PtdSer decarboxylation, even in the presence of ethanolamine (22, 23). In all these studies difficulties in measurement of pool sizes and possible non-homogeneous radiolabeling of lipid precursor pools complicate interpretation of the data. Several cell types, for example rat mammary carcinoma cells (24), hybridoma cells (25), and human keratinocytes (26), have an absolute requirement for ethanolamine for growth and/or proliferation (27). One cannot, however, conclude that the ethanolamine requirement of these cells is necessarily for PtdEtn synthesis, since ethanolamine is also used for the glycosylphosphatidylinositol anchors of proteins (28).

Little information is available on how PtdSer biosynthesis is regulated and whether or not the two pathways for PtdEtn synthesis are coordinately regulated. We have therefore cloned and expressed a cDNA that encodes mouse liver PtdSer synthase-1. First, we demonstrate that this cDNA complements the ethanolamine auxotrophy of M.9.1.1 mutant CHO-K1 cells. We also show that when PtdSer synthesis is increased severalfold in hepatoma cells overexpressing mouse liver PtdSer synthase-1 cDNA, PtdSer and PtdEtn levels remain the same as in control cells. Apparently, as compensation for overexpression of PtdSer synthase-1 activity, the catabolism of PtdSer and/or PtdSer-derived PtdEtn is increased, and the CDP-ethanolamine pathway for PtdEtn biosynthesis is inhibited.

    EXPERIMENTAL PROCEDURES
Top
Abstract
Introduction
Procedures
Results
Discussion
References

Materials-- CHO-K1 cells and McArdle 7777 cells were obtained from the American Type Tissue Culture Collection. The radiochemicals [3-3H]serine, [1-3H]ethanolamine, [methyl14C]choline, and [32P]ATP were from Amersham Pharmacia Biotech (Oakville, Ontario, Canada). Cytidine-5'-diphospho-[1,2-14C]ethanolamine was from ICN Radiochemicals (Montreal, Quebec, Canada). Thin layer chromatography plates were purchased from BDH Chemicals. Fetal bovine serum, horse serum, tissue culture media, and DNA modifying enzymes were purchased from Life Technologies, Inc. Authentic phospholipid standards were from Avanti Polar Lipids (Birmingham, AL). Sodium orthovanadate was purchased from Calbiochem. The phosphatase 1/2A inhibitor, microcystin, was a generous gift from Dr. C. Holmes, University of Alberta. All other chemicals were from Sigma or Fisher.

Oligonucleotides were synthesized as primers for PCR. Modified lambda gt11 forward (FP) and reverse (RP) sequencing primers, and two gene-specific primers based on the CHO-K1 PtdSer synthase-1 cDNA sequence (10), were synthesized. The oligonucleotide 520S was complementary to the sense strand of PtdSer synthase-1, and 913AS was complementary to the antisense strand (FP, GCGACGACTCCTGGAGCCCG; RP, TGACACCAGACCAACTGGTAATG; 520S, GGCCATGAAGGCCTTGTTGATCCGTAGT; 913AS, TATGAATGTCCTTGAAGCTTGCCCA).

PCR and Cloning-- The cDNA from a lambda gt11 mouse liver expression library (CLONTECH, Palo Alto, CA) was isolated by modification of the method described by Sambrook et al. (29). Luria-Bertani medium (500 ml) was inoculated with Y1090R- strain Escherichia coli infected with lambda  phage from the mouse liver cDNA library. Cells were grown at 37 °C until lysis was apparent, after which addition of 10 ml of chloroform and further incubation at 37 °C for 10 min completed lysis. DNase I and RNase A were added to the lysed cultures at final concentrations of 1 and 5 µg/ml, respectively, and incubated at 37 °C for 30 min. Polyethylene glycol and NaCl were added at final concentrations of 10% and 1 M, respectively. Cultures were maintained at 4 °C for 1 h to precipitate the phage, which were isolated by centrifugation for 10 min at 10,000 × g. The phage-containing pellet was resuspended in 10 mM Tris-HCl buffer (pH 8.0) containing 10 mM EDTA and 0.5% SDS, followed by incubation at 68 °C for 20 min to release the lambda  phage cDNA. The DNA solution was extracted three times with an equal volume of phenol:chloroform (1:1), and the DNA was precipitated by addition of NaCl (to a final concentration of 0.25 M) and 2 volumes of 95% ethanol. The DNA was pelleted by centrifugation for 10 min at 10,000 × g. The pellet was washed with 80% ethanol, and the DNA was resuspended at a final concentration of 1 µg/µl in 10 mM Tris-HCl buffer (pH 8.0) containing 1 mM EDTA and used as a template for PCR reactions. The PCR reaction mixture (50 µl) contained 25 ng of lambda  phage cDNA, 5 µl of the supplied 10-fold concentrated reaction buffer (Panvera, Madison, WI), 2.5 mM MgCl, 0.25 mM of each nucleotide triphosphate, 10 pmol of lambda gt11 sequencing primer (FP or RP), and 10 pmol of PtdSer synthase-1-specific primers (520S or 913AS), as well as 2.5 units of TaKaRa Ex Taq DNA polymerase (Panvera, Madison, WI). The PCR reaction was performed for 30 cycles at 94 °C for 1 min, 65 °C for 2 min, 72 °C for 1.5 min, and a final 10-min extension at 72 °C. The products of the PCR reaction were separated by agarose gel electrophoresis. The DNA fragments were eluted using a gel extraction kit (Qiagen, Missisauga, Ontario, Canada) and cloned into the pCRII vector (Invitrogen, San Diego, CA). Inserts were sequenced using the dideoxy termination/polyacrylamide gel method (30) by the DNA Core facility, at the University of Alberta. The full-length clone was sequenced in both directions.

Construction of a Full-length PtdSer Synthase-1 cDNA and Expression of Mouse Liver PtdSer Synthase-1 cDNA in Mammalian Cells-- The 950-bp 5' end generated by 913AS/FP primers was ligated into the KpnI/HindIII sites of pBluescript (SK+) (Stratagene). The 1840-bp 3' end generated by 520S/RP primers was inserted into the ApaI/SpeI sites of pBluescript (SK+). The 3' end was then digested with HindIII/NotI and joined in-frame to the 5' fragment. A 2-kilobase pair fragment containing the entire coding region of PtdSer synthase-1 was inserted into the eukaryotic expression vector pRc/CMV (Invitrogen).

M.9.1.1 cells (14) (a gift from Dr. D. R. Voelker, National Jewish Research Center, Denver, CO) and McArdle 7777 rat hepatoma cells were grown in a 5% CO2 incubator at 37 °C. M.9.1.1 cells were routinely maintained in Ham's F-12 medium containing 10% delipidated fetal bovine serum (31) and 20 µM ethanolamine. McArdle 7777 cells were maintained in Dulbecco's modified Eagle's medium containing 10% fetal bovine serum and 10% horse serum. Both types of cells were transfected by the calcium phosphate precipitation method (32) using 10 µg of DNA. For selection of stable transfectants, cells were cultured in medium containing 400 µg/ml G418. Individual colonies were isolated, and once cell lines were established the concentration of G418 was reduced to 200 µg/ml. The control cells for the experiments with M.9.1.1 and McArdle 7777 were transfected with vector alone.

Enzyme Activities-- PtdSer synthase activity was measured as described previously (33). Briefly, cells were scraped from dishes and disrupted by sonication with a probe sonicator, 2 × 10 s, in 10 mM HEPES buffer (pH 7.5) containing 0.25 M sucrose. Lysates were centrifuged for 2 min at 600 × g to pellet cellular debris, and PtdSer synthase-1 activity was measured in the supernatant using [3-3H]serine, [1-3H]ethanolamine, or [methyl-14C]choline. The activities of the three enzymes of the CDP-ethanolamine pathway were determined as described previously (33-35). The products of the ethanolamine kinase and ET assays were separated by thin layer chromatography on silica gel G-60 thin layer plates in the solvent system methanol, 0.6% NaCl, NH4OH (10:10:1, v/v) and identified by comparison to authentic standards. In some experiments, as indicated, ET was assayed in lysates that had been prepared in the presence of the phosphoprotein phosphatase inhibitors microcystin (5 µM) (36), sodium orthovanadate (1 mM), and sodium fluoride (10 mM).

Radioabeling of Cells-- PtdSer and PtdEtn were radiolabeled by incubation of cultured cells with [3-3H]serine (5 µCi/ml). For experiments in which cells were labeled with [1-3H]ethanolamine, radioactivity in PtdEtn was determined. In addition, water-soluble intermediates of the CDP-ethanolamine pathway were separated by thin layer chromatography on silica gel G-60 thin layer plates in a combination of two different solvent systems. Ethanolamine and phosphoethanolamine were separated in the solvent system methanol, 0.6% NaCl, NH4OH (10:10:1, v/v), whereas CDP-ethanolamine and glycerophosphoethanolamine were separated in the solvent system ethanol, 0.9% NaCl, NH4OH (80:10:26, v/v). Ethanolamine, phosphoethanolamine, CDP-ethanolamine, and glycerophosphoethanolamine standards were added as carriers. The bands corresponding to these intermediates were scraped, and radioactivity was measured.

Other Analyses-- Phospholipids were extracted from cells by the method of Bligh and Dyer (37) and separated by thin layer chromatography on silica gel G-60 thin layer chromatography plates in the solvent chloroform:methanol:acetic acid:formic acid:water (70:30:12:4:2, v/v). Phospholipids were visualized by exposure to iodine vapors and identified by comparison with authentic standards. The phosphorus content of each phospholipid was determined as described previously (38). For measurement of the cellular content of diacylglycerols, lipids were separated by thin layer chromatography in the solvent system heptane:isopropyl ether:acetic acid (60:40:4, v/v). The thin layer plate was immersed in a solution of cupric acetate/phosphoric acid and then heated at 180 °C for 5 min to visualize the lipids (39). The amount of diacylglycerol was determined by densitometric scanning of the bands and comparison with known amounts of standard diacylglycerol. The triacylglycerol content of the cells was determined in a total lipid extract using the Triglyceride E Kit (Wako Chemicals, Richmond, VA). Protein concentrations were determined by the BCA method (Pierce).

    RESULTS
Top
Abstract
Introduction
Procedures
Results
Discussion
References

Isolation of the cDNA for Mouse Liver PtdSer Synthase-1-- PCR cloning was used to isolate the cDNA for PtdSer synthase-1 from mouse liver. cDNA from a lambda gt11 cDNA library containing 2 × 106 clones was isolated and used as a template for PCR reactions. The 5' and 3' ends of PtdSer synthase-1 were amplified in two separate PCR reactions. One fragment, 520S/RP, contained the 3' end, and the other fragment, 913AS/FP, contained the 5' end. These two fragments possessed a 235-bp overlapping region and were ligated at a common HindIII restriction site. The assembled PtdSer synthase-1 clone was 2362 bp in length with the longest open reading frame encoding a 473-amino acid protein with a predicted molecular weight of 55,617 (Fig. 2). The 5'-untranslated region contained an in-frame stop codon suggesting that the clone was full length.


View larger version (85K):
[in this window]
[in a new window]
 
Fig. 2.   Nucleotide and predicted amino acid sequences of mouse liver PtdSer synthase-1 cDNA. The translational start codon is numbered +1. The first ATG and the stop codon of the largest open reading frame are underlined. An in-frame stop codon (TAG) in the 5'-untranslated region is in boldface. Five putative transmembrane regions are indicated by broken underlines.

The PtdSer synthase-1 sequence lacks a typical N-terminal signal sequence for targeting the protein to the ER but contains five putative membrane spanning regions according to hydropathy plot analysis (40). In addition, the C terminus contains the Lys-Lys motif that has been proposed to be an ER membrane retention signal (41). These findings, as well as the observation that immunoreactive PtdSer synthase-1 protein is present in the ER of CHO cells (11), are consistent with the enzymatic activity of PtdSer synthase-1 being primarily localized to the ER (33). A human myeloblast cDNA sequence encoding a putative PtdSer synthase-1 was identified by sequence analysis from GenBankTM (accession number D14694). Comparison of the PtdSer synthase-1 sequences of mouse liver, CHO-K1 (GenBankTM accession number D90468), and human myeloblast cells revealed a very high degree of conservation across species (Fig. 3). However, the mouse liver PtdSer synthase-1 cDNA was not homologous to the PtdSer synthase cDNAs isolated from E. coli, S. cerevisiae, or Bacillus subtilis (7, 8, 42, 43). This lack of homology was not unexpected since the yeast and bacterial enzymes synthesize PtdSer by a reaction different from that in mammalian cells and use CDP-diacylglycerol as a substrate instead of catalyzing a base-exchange reaction. At the level of their cDNAs, the three mammalian clones are ~80% identical, whereas their amino acid sequences are >90% identical (Fig. 3). In contrast, only 30% of the amino acids in the mouse liver PtdSer synthase-1 are identical to those deduced from the PtdSer synthase-2 cDNA which has recently been cloned from CHO-K1 cells (13).


View larger version (69K):
[in this window]
[in a new window]
 
Fig. 3.   Comparison of the predicted amino acid sequences of PtdSer synthases-1 from CHO-K1 cells, human myeloblastic cells, and mouse liver. Sequences of PtdSer synthase-1 from CHO-K1 (10) and human myeloblastic cells (GenBankTM accession number D14694) were aligned with the deduced amino acid sequence of mouse liver PtdSer synthase-1. Identical amino acids are enclosed in boxes.

Complementation of the CHO-K1 Mutant Cells, M.9.1.1-- To confirm that the isolated mouse liver cDNA encoded PtdSer synthase-1, we first determined whether or not the cDNA complemented the growth defect of M.9.1.1 cells and restored the phospholipid composition of these cells. Voelker and Frazier (14) have reported that PtdSer synthase activity in M.9.1.1 cells is 50% lower than in wild-type CHO-K1 cells, an observation that our data confirm (Fig. 4). Stable expression of PtdSer synthase-1 cDNA in M.9.1.1 cells resulted in an approximately 5-fold increase in the incorporation of [3-3H]serine into PtdSer in a cell-free assay (Fig. 4). In addition, PtdSer synthase-1 utilized both ethanolamine and choline as substrates for base-exchange reactions. Compared with control M.9.1.1 cells that were transfected with vector alone, a 4-fold increase in base exchange activity for the incorporation of [1-3H]ethanolamine into PtdEtn and a 6-fold increase in the in vitro incorporation of [methyl-14C]choline into PtdCho were observed in M.9.1.1 cells expressing mouse liver PtdSer synthase-1 (Fig. 4). Since the cloned mouse liver PtdSer synthase-1 increased the choline exchange activity, the isolated cDNA does not encode PtdSer synthase-2 which lacks this activity (13).


View larger version (23K):
[in this window]
[in a new window]
 
Fig. 4.   Base exchange enzyme activities of wild-type CHO-K1 cells, M.9.1.1 cells, and control cells expressing mouse liver PtdSer synthase-1 (M.9.1.1/MLPSS1). Base exchange activities were measured using [3-3H]serine (Ser), [1-3H]ethanolamine (Etn), and [methyl-14C]choline (Cho) in cellular lysates from CHO-K1 cells (solid bars), control M.9.1.1 cells transfected with vector alone (open bars), and M.9.1.1/MLPSS1 cells (stippled bars). Data are averages ± S.D. of triplicate analyses from one experiment that was representative of three similar experiments.

M.9.1.1 cells are defective in the synthesis of PtdSer and, consequently, in the production of PtdEtn from PtdSer decarboxylation. In these cells ethanolamine (20 µM) is required in the culture medium to support sufficient PtdEtn synthesis via the CDP-ethanolamine pathway for normal growth (14). We therefore examined whether or not M.9.1.1 cells could be transformed into ethanolamine prototrophs by expression of mouse liver PtdSer synthase-1 cDNA. As shown in Fig. 5, and as previously reported (14), M.9.1.1 cells grew in the absence of ethanolamine for approximately three generations, then died. Addition of 20 µM ethanolamine restored normal growth. However, M.9.1.1 cells expressing PtdSer synthase-1 grew at the same rate regardless of the presence of ethanolamine (Fig. 5). Thus, expression of PtdSer synthase-1 cDNA complemented the ethanolamine auxotrophy of M.9.1.1 cells.


View larger version (16K):
[in this window]
[in a new window]
 
Fig. 5.   Growth of M.9.1.1 cells and M.9.1.1 cells expressing mouse liver PtdSer synthase-1 (M.9.1.1/MLPSS1 cells). Control M.9.1.1 cells transfected with vector alone (squares) and M.9.1.1/MLPSS1 cells (circles) were plated at a density of 5 × 104 cells/60-mm dish and cultured in Ham's F-12 medium supplemented with 10% delipidated fetal bovine serum with (open symbols) or without (solid symbols) 20 µM ethanolamine. Cells were harvested at 24-h intervals by trypsinization and counted. Data are averages ± S.D. of triplicate analyses from one representative experiment of three similar experiments. Some error bars are hidden by the symbols.

Analysis of the phospholipid composition of M.9.1.1 cells revealed that PtdSer and PtdEtn levels were significantly (57 and 51%, respectively (p < 0.05)) lower when the cells were cultured in the absence of ethanolamine than in the presence of ethanolamine (Fig. 6), in agreement with previous data (14). However, the content of PtdSer and PtdEtn in M.9.1.1 cells expressing mouse liver PtdSer synthase-1 cDNA, cultured in either the presence or absence of exogenous ethanolamine, was very similar to that of M.91.1 cells grown in the presence of 20 µM ethanolamine (Fig. 6). These results are consistent with those from studies by Kuge et al. (10) in which expression of PtdSer synthase-1 cDNA from CHO-K1 cells corrected the growth defect and normalized the phospholipid composition of PSA-3 cells.


View larger version (12K):
[in this window]
[in a new window]
 
Fig. 6.   PtdSer and PtdEtn content of control M.9.1.1 cells and M.9.1.1 cells expressing mouse liver PtdSer synthase-1. Cells were grown in medium supplemented with 20 µM ethanolamine for 48 h. Medium with (+) or without (-) 20 µM ethanolamine (Eth) was then added for an additional 48 h after which cells were harvested, and the content of PtdSer and PtdEtn was determined. PSS (+) denotes M.9.1.1 cells transfected with PtdSer synthase-1 cDNA; PSS (-) denotes control M.9.1.1 cells transfected with vector alone. Data are averages ± S.D. of triplicate analyses from three independent experiments. *, statistical significance (p < 0.05) of differences between cells cultured in the presence and absence of 20 µM ethanolamine was evaluated by the Student's t test.

Expression of Mouse Liver PtdSer Synthase-1 in McArdle 7777 Cells-- We next expressed PtdSer synthase-1 cDNA in McArdle 7777 rat hepatoma cells that possess endogenous PtdSer synthase-1 activity. Several stably transfected McArdle 7777 cell lines expressing mouse liver PtdSer synthase-1 were generated. We selected one cell line (designated Mc/PSS1) that exhibited a 2-3-fold increase in serine, ethanolamine, and choline exchange activities when the cell lysate was assayed in vitro (Fig. 7). Although we screened about 50 transfectants, a 3-fold increase in PtdSer synthase-1 activity was the maximum achieved. One explanation for this observation is that a higher expression of PtdSer synthase-1 might be lethal. Mc/PSS1 cells and control cells (i.e. McArdle 7777 cells transfected with vector alone) were labeled with [3-3H]serine for up to 1 h, and incorporation of radiolabel into PtdSer and PtdEtn was determined. The total uptake of [3H]serine was the same in the two cell types. In Mc/PSS1 cells the rate of incorporation of [3H]serine into PtdSer was 3-fold higher than in control cells, and the rate of incorporation into PtdEtn was double that of control cells (Fig. 8). These data indicate that the synthesis of both PtdSer and PtdEtn was increased in Mc/PSS1 cells compared with control cells. Fig. 9 shows that the amounts of the major phospholipids were not significantly different in Mc/PSS1 cells and control cells.


View larger version (11K):
[in this window]
[in a new window]
 
Fig. 7.   Base exchange activities of control McArdle 7777 cells and McArdle 7777 cells expressing mouse liver PtdSer synthase-1 (Mc/PSS1). Base exchange activities were measured in cellular lysates using [3-3H]serine (Ser), [methyl-14C]choline (Cho), and [1-3H]ethanolamine (Etn). Solid symbols, control McArdle 7777 cells transfected with vector alone; open symbols, Mc/PSS1 cells. Data are averages ± S.D. of triplicate analyses from three independent experiments.


View larger version (19K):
[in this window]
[in a new window]
 
Fig. 8.   Incorporation of [3H]serine into PtdSer and PtdEtn of control cells and Mc/PSS1 cells. Cells were incubated in medium containing [3-3H]serine (20 µCi/dish). At the indicated times, cells were harvested, and PtdSer and PtdEtn were extracted and separated by thin layer chromatography. Open symbols, control McArdle cells transfected with vector alone; solid symbols, Mc/PSS1 cells. Data are averages ± S.D. of triplicate analyses from one experiment which was repeated twice with similar results. Some error bars are obscured by the symbols.


View larger version (12K):
[in this window]
[in a new window]
 
Fig. 9.   Phospholipid composition of control cells and Mc/PSS1 cells. Cells were grown to 80% confluence and then were harvested, and the phospholipids were extracted and separated by thin layer chromatography. The phosphorous content of PtdSer, PtdEtn, PtdCho, sphingomyelin (SM), and phosphatidylinositol (PtdIns) was determined. Solid symbols, control McArdle 7777 cells transfected with vector alone; open symbols, Mc/PSS1 cells. Data are averages ± S.D. of triplicate analyses from three independent experiments.

These observations suggested that the levels of PtdSer and PtdEtn might be tightly controlled and that synthesis and/or degradation of these lipids might be regulated. We therefore examined further the metabolism of PtdSer and PtdEtn in control cells and Mc/PSS1 cells. The cells were pulse-labeled for 1 h with [3-3H]serine, then the radiolabel was chased for up to 12 h (Fig. 10). As expected from the data shown in Fig. 8, incorporation of [3H]serine into PtdSer in Mc/PSS1 cells at the end of the 1-h pulse was approximately 2.5-fold higher than in control cells (Fig. 10, upper panel). However, radioactivity in PtdSer declined more rapidly in Mc/PSS1 cells than in control cells, indicating an increased rate of degradation of PtdSer in the cells overexpressing PtdSer synthase-1. Fig. 10 (lower panel) also shows that the incorporation of [3H]serine into PtdEtn was consistently higher in Mc/PSS1 cells than in control cells, suggesting that at least a portion of the increased radiolabel lost from PtdSer in Mc/PSS1 cells was due to an increased conversion of PtdSer to PtdEtn. However, in neither Mc/PSS1 nor control cells was the radioactivity lost from PtdSer quantitatively recovered in PtdEtn. For example, in Mc/PSS1 cells during the first 6 h of the chase period, 107 × 103 dpm/mg protein were lost from PtdSer, whereas the 3H content of PtdEtn increased by only 25 × 103 dpm/mg protein. Similarly, during the same period, 35 × 103 dpm/mg protein were lost from PtdSer of control cells, and only 15 × 103 dpm/mg protein accumulated in PtdEtn (Fig. 10). These data show the following: (i) in both types of cells some PtdSer was converted into PtdEtn; (ii) the conversion of PtdSer to PtdEtn was increased in Mc/PSS1 cells compared with control cells; (iii) some radiolabeled PtdSer and/or PtdEtn was degraded in both Mc/PSS1 and control cells; and (iv) in Mc/PSS1 cells, ~four times as much radiolabeled PtdSer and/or PtdEtn was degraded as in control cells. Presumably, these metabolic events are coordinated to ensure constant cellular levels of PtdSer and PtdEtn.


View larger version (20K):
[in this window]
[in a new window]
 
Fig. 10.   Metabolism of [3H]serine-derived PtdSer and PtdEtn in control cells and Mc/PSS1 cells. Cells were incubated for 1 h in medium containing [3H]serine (20 µCi/dish). The medium was then removed, and fresh medium containing 1 mM unlabeled serine was added. At the indicated times cells were harvested, and PtdSer (upper panel) and PtdEtn (lower panel) were extracted and separated by thin layer chromatography. Open symbols, control McArdle 7777 cells transfected with vector alone; solid symbols, Mc/PSS1 cells. Data are averages ± S.D. from triplicate analyses from one experiment which was repeated at twice with similar results. Some error bars are too small to be visible.

Overexpression of PtdSer Synthase-1 Inhibits PtdEtn Synthesis via the CDP-ethanolamine Pathway-- Figs. 8 and 10 show that Mc/PSS1 cells produced more PtdEtn by decarboxylation of PtdSer than did control cells. However, the total cellular mass of PtdEtn did not increase (Fig. 9). We therefore examined whether or not the increased expression of mouse liver PtdSer synthase-1 in McArdle 7777 cells inhibited the synthesis of PtdEtn via the other major route for PtdEtn biosynthesis, the CDP-ethanolamine pathway. Mc/PSS1 cells were incubated with [1-3H]ethanolamine for up to 1 h, and incorporation of radioactivity was measured in PtdEtn and intermediates of the CDP-ethanolamine pathway (Fig. 11). The total amount of [3H]ethanolamine taken up by control and Mc/PSS1 cells was the same. In Mc/PSS1 cells, more radiolabeled ethanolamine, phosphoethanolamine, and CDP-ethanolamine was present than in control cells. In contrast, the incorporation of radiolabel into PtdEtn was greatly reduced in Mc/PSS1 cells compared with control cells. The relative pool sizes of radiolabeled ethanolamine metabolites in Mc/PSS1 cells and control cells were also compared after labeling with [3H]ethanolamine for 24 h. As expected from the data shown in Fig. 11, more radiolabeled ethanolamine and phosphoethanolamine (2.5- and 13-fold, respectively) were present in Mc/PSS1 cells than in control cells. In control cells, the dpm/mg protein in ethanolamine were (3.30 ± 0.50) × 103, and in Mc/PSS1 cells the dpm/mg protein were (8.20 ± 2.50) × 103. In phosphoethanolamine the dpm/mg protein were (6.60 ± 2.50) × 103 in control cells and (84.7 ± 20.5) × 103 in Mc/PSS1 cells. For CDP-ethanolamine, (2.10 ± 0.20) × 103 dpm/mg protein were recovered from control cells and (3.40 ± 1.40) × 103 dpm/mg protein from Mc/PSS1 cells.


View larger version (22K):
[in this window]
[in a new window]
 
Fig. 11.   Incorporation of [3H]ethanolamine into PtdEtn and ethanolamine-derived metabolites in control McArdle 7777 cells and Mc/PSS1 cells. Cells were incubated in medium containing [3H]ethanolamine (1.5 µCi/dish) for the indicated times and then harvested, and PtdEtn and water-soluble intermediates of the CDP-ethanolamine pathway (Etn, ethanolamine; Etn-P, phosphoethanolamine; CDP-Etn, CDP-ethanolamine) were extracted and separated by thin layer chromatography. Open symbols, control McArdle 7777 cells transfected with vector alone; solid symbols, Mc/PSS1 cells. Data are averages ± S.D. of triplicate analyses from one experiment which was repeated twice with similar results. Some error bars are obscured by symbols.

Two likely explanations for the decreased labeling of PtdEtn from [3H]ethanolamine when PtdSer synthase-1 was overexpressed are as follows: (i) PtdEtn synthesis via the CDP-ethanolamine pathway was inhibited, or (ii) the rate of degradation of ethanolamine-derived PtdEtn was increased. These two possibilities were distinguished by incubation of Mc/PSS1 cells with [3H]ethanolamine for 24 h. The radioactivity was chased with 2 mM unlabeled ethanolamine, and the amounts of [3H]PtdEtn and ethanolamine-derived intermediates were measured. Fig. 12 shows that the rate of loss of radiolabel from PtdEtn was the same in Mc/PSS1 cells as in control cells. In addition, the amounts of labeled degradation products of PtdEtn (i.e. [3H]phosphoethanolamine, [3H]ethanolamine, and [3H]glycerophosphoethanolamine) were the same in Mc/PSS1 and control cells (data not shown). The combined data, therefore, indicate that the increased content of [3H]ethanolamine-labeled intermediates in Mc/PSS1 cells, as shown in Fig. 11, was due to inhibition of PtdEtn synthesis via the CDP-ethanolamine pathway rather than increased degradation of ethanolamine-derived PtdEtn.


View larger version (14K):
[in this window]
[in a new window]
 
Fig. 12.   Degradation of [3H]ethanolamine-derived PtdEtn. Control McArdle 7777 cells transfected with vector alone (closed symbols) and Mc/PSS1 cells (open symbols) were labeled with [3H]ethanolamine (1.5 µCi/dish) for 24 h. The radiolabeled medium was removed and replaced with medium containing (2 mM) unlabeled ethanolamine. Cells were harvested at the indicated times, and incorporation of radiolabel into PtdEtn was determined. Data are averages ± S.D. of triplicate analyses from one experiment which was repeated twice with similar results.

Since in Mc/PSS1 cells less PtdEtn was synthesized by the CDP-ethanolamine pathway than in control cells, we considered the possibility that the activity of one of the enzymes of this pathway, most likely ET, was decreased in Mc/PSS1 cells. ET has been suggested to be the rate-limiting enzyme of the CDP-ethanolamine pathway under most metabolic conditions (44-46). However, Table I shows that the activities of ethanolamine kinase, ET, and ethanolaminephosphotransferase in Mc/PSS1 cells were essentially the same as in control cells. Since many enzymes are regulated by phosphorylation-dephosphorylation events, ET was also assayed in cell lysates that had been prepared in the presence of the phosphoprotein phosphatase inhibitors microcystin (36), vanadate, and fluoride, and the same results were obtained.

                              
View this table:
[in this window]
[in a new window]
 
Table I
Activities of enzymes of the CDP-ethanolamine pathway in control cells and Mc/PSS1 cells
Total cellular membranes and cytosol were prepared by centrifugation of the cell lysate at 400,000 × g for 30 min. Cytosol was assayed for ethanolamine kinase and ET activity. Membranes were assayed for CDP-ethanolamine:1,2-diacylglycerol ethanolaminephosphotransferase activity. The data for each enzyme assay are from triplicate analyses of one experiment which was repeated twice with similar results. ET activity was assayed in the presence (+) or absence (-) of 5 µM microcystin, 1 mM sodium orthovanadate, and 10 mM NaF.

The level of diacylglycerol has also been reported to regulate the CDP-ethanolamine pathway (47, 48) since this lipid is a substrate for ethanolaminephosphotransferase. We therefore measured the diacylglycerol content of control and Mc/PSS1 cells and found that the amount of this lipid was approximately the same in control cells (4.4 ± 0.5 nmol/mg protein) and (4.0 ± 0.5 nmol/mg protein) Mc/PSS1 cells. Since diacylglycerol is an intermediate in the biosynthesis of several important glycerolipids and has also been implicated as a lipid second messenger, the cellular content of this lipid probably fluctuates only transiently and locally. Consequently, changes in the supply of diacylglycerols for ethanolaminephosphotransferase might not be detected by measurement of the total cellular diacylglycerol content. We therefore measured the cellular content of triacylglycerols as a potential "reserve" precursor pool of diacylglycerols. The content of triacylglycerols was 40% less in Mc/PSS1 cells overexpressing PtdSer synthase-1 (14.2 ± 1.2 nmol/mg protein) than in control cells (23.7 ± 2.1 nmol/mg protein).

    DISCUSSION
Top
Abstract
Introduction
Procedures
Results
Discussion
References

We report the cloning and expression of a cDNA that encodes PtdSer synthase-1 from mouse liver and show that the PtdSer synthesis and/or decarboxylation pathways can coordinately regulate the production of PtdEtn via the CDP-ethanolamine pathway.

As evidence that the cloned cDNA encoded PtdSer synthase-1, the cDNA for mouse liver PtdSer synthase-1 was heterologously expressed in M.9.1.1 cells that are mutant CHO cells deficient in PtdSer synthase-1 activity (14). When PtdSer synthase-1 cDNA was expressed, the ethanolamine auxotrophy of the M.9.1.1 cells was eliminated, the base exchange activities with serine, choline, and ethanolamine were increased, and the levels of PtdSer and PtdEtn were restored to normal. In addition, expression of mouse liver PtdSer synthase-1 in McArdle 7777 rat hepatoma cells resulted in increased in vitro base exchange activity and an increased rate of incorporation of [3H]serine into PtdSer and PtdEtn in intact cells. These observations are consistent with previous data showing that a PtdSer synthase-1 cDNA from CHO-K1 cells complemented the ethanolamine auxotrophy of a mutant CHO cell line, PSA-3 (10).

Our studies in McArdle hepatoma cells imply that the metabolism of PtdSer and PtdEtn is coordinately regulated so that cellular levels of these lipids remain constant. When PtdSer synthase-1 activity was overexpressed, homeostasis of PtdSer and PtdEtn was maintained by the following: (i) increased conversion of PtdSer to PtdEtn, (ii) increased degradation of PtdSer and/or PtdEtn derived from PtdSer, and (iii) inhibition of the CDP-ethanolamine pathway for PtdEtn synthesis. These findings are reminiscent of those from a study in which CTP:phosphocholine cytidylyltransferase, the rate-limiting enzyme of PtdCho biosynthesis via the CDP-choline pathway, was overexpressed in COS cells (49). Despite a greatly increased rate of PtdCho synthesis in these transfected cells, the mass of PtdCho was barely increased. However, the rate of PtdCho degradation was increased, presumably as a mechanism for maintaining a constant level of PtdCho. Similarly, overexpression of E. coli PtdSer synthase in E. coli did not alter the phospholipid composition of these cells (50).

The increased conversion of PtdSer to PtdEtn in cells overexpressing PtdSer synthase is most likely the direct result of increased PtdSer synthesis. Production of PtdSer-derived PtdEtn appears not to be limited by the activity of the decarboxylase but rather by availability of the substrate, PtdSer (51). PtdSer supply is, in turn, regulated by either the rate of PtdSer synthesis or the rate of translocation of PtdSer from its site of synthesis on the ER to the site of the decarboxylase on the outer aspect of inner mitochondrial membranes (15). Hence, an increased supply of PtdSer to mitochondria would be expected to be translated into an increased production of PtdEtn by the decarboxylase.

A second consequence of overexpression of PtdSer synthase-1 was that degradation of PtdSer and/or PtdSer-derived PtdEtn was increased, although the degradation of PtdEtn derived from the CDP-ethanolamine pathway was not increased. Most likely the enhanced degradation was of PtdSer rather than of PtdEtn since in primary hepatocytes newly made PtdSer is rapidly degraded, whereas PtdSer-derived PtdEtn is not significantly degraded (52).

A third consequence of overexpression of PtdSer synthase-1 in McArdle 7777 cells was that the synthesis of PtdEtn via the CDP-ethanolamine pathway was inhibited, as was demonstrated by the greatly reduced incorporation of [3H]ethanolamine into PtdEtn. This reduction in incorporation of radioactivity could not be accounted for by an increased degradation of ethanolamine-derived PtdEtn. Moreover, the amounts of [3H]ethanolamine-labeled precursors of PtdEtn were increased indicating that the CDP-ethanolamine pathway for PtdEtn synthesis was inhibited. We therefore investigated which step of the CDP-ethanolamine pathway was modified when PtdSer synthase-1 was overexpressed. Under most metabolic conditions the rate-limiting enzyme of this pathway is thought to be ET (44-46). However, the total in vitro activity of ET was unaffected by overexpression of PtdSer synthase-1. We cannot, however, exclude the possibility that the distribution of ET between a small active pool and a large inactive pool had been altered since such a change would not have been detected in our experiments. Unlike the cytidylyltransferase of the CDP-choline biosynthetic pathway, there is no evidence that the activity of ET is regulated by reversible translocation to and from membranes (44). ET is presumed to be a cytosolic protein that neither requires lipids for activity nor is tightly associated with membranes (47, 53-55). However, some association of ET with ER membranes has been detected in immunogold electron microscopy studies (53). The cDNA sequences of ET (56) and CTP:phosphocholine cytidylyltransferase (57) share some similarities, especially at the N termini of the corresponding proteins. CTP:phosphocholine cytidylyltransferase contains an amphipathic alpha -helical C-terminal domain that has been proposed to mediate membrane association (58). From analysis of the cDNA sequence of ET (56) it appears that ET might also have an amphipathic alpha -helical domain close to the C terminus that could potentially form a loose association with membranes.

The supply of diacylglycerols has also been implicated in the regulation of the CDP-ethanolamine pathway (47, 48) by virtue of this lipid being a substrate for ethanolaminephosphotransferase. Similarly, under some metabolic conditions, diacylglycerols can regulate the CDP-choline pathway for PtdCho synthesis at the level of the cholinephosphotransferase (59-61). We detected no difference in the diacylglycerol content of control cells and Mc/PSS1 cells although our experiments would not have detected a change in a small localized pool of this lipid. However, in cells overexpressing PtdSer synthase-1, the cellular content of triacylglycerols was 40% less than in control cells. For each molecule of PtdSer synthesized, one molecule of phospholipid, and hence one molecule of diacylglycerol, is consumed. If the "reservoir" of stored triacylglycerols in the cell served as a precursor pool of diacylglycerols, a reduction in triacylglycerol mass might result from the increased demand for diacylglycerols in the cells overexpressing PtdSer synthase-1. Therefore, overexpression of PtdSer synthase-1 might deplete a specific pool of diacylglycerols that is used for PtdEtn synthesis via the ethanolaminephosphotransferase reaction.

In conclusion, an increased synthesis of PtdSer, induced by overexpression of mouse liver PtdSer synthase-1 cDNA, increases the production of PtdEtn from the PtdSer decarboxylation pathway and inhibits the synthesis of PtdEtn via the CDP-ethanolamine pathway. These observations suggest that the synthesis and decarboxylation of PtdSer, as well as the synthesis of PtdEtn from the CDP-ethanolamine pathway and the degradation of PtdSer and/or PtdEtn, are coordinately regulated so that constant cellular levels of these phospholipids are maintained. Experiments are presently underway in our laboratory to determine whether or not increasing in the synthesis of PtdEtn via the CDP-ethanolamine pathway, by overexpression of ET, reciprocally inhibits the synthesis and/or decarboxylation of PtdSer.

    FOOTNOTES

* The costs of publication of this article were defrayed in part by the payment of page charges. The article must therefore be hereby marked "advertisement" in accordance with 18 U.S.C. Section 1734 solely to indicate this fact.

The nucleotide sequence(s) reported in this paper has been submitted to the GenBankTM/EMBL Data Bank with accession number(s) AF042731.

To whom correspondence should be addressed: Lipid and Lipoprotein Research Group, 315 Heritage Medical Research Center, University of Alberta, Edmonton, AB T6G 2S2, Canada. Tel.: 403-492-7250; Fax: 403-492-3383; E-mail: jean.vance{at}ualberta.ca.

1 The abbreviations used are: PtdSer, phosphatidylserine; CHO-K1, Chinese hamster ovary; ER, endoplasmic reticulum; ET, CTP:ethanolamine-phosphate cytidylyltransferase; PtdCho, phosphatidylcholine; PtdEtn, phosphatidylethanolamine; bp, base pair(s); PCR, polymerase chain reaction.

    REFERENCES
Top
Abstract
Introduction
Procedures
Results
Discussion
References

  1. Nishizuka, Y. (1992) Science 258, 607-614[Medline] [Order article via Infotrieve]
  2. Schroit, A. J., and Zwaal, R. F. A. (1991) Biochim. Biophys. Acta 1071, 313-329[Medline] [Order article via Infotrieve]
  3. Bevers, E., Comfurius, P., van Rijn, J., Hemker, H., Zwaal, R. (1982) Eur. J. Biochem. 122, 429-436[Medline] [Order article via Infotrieve]
  4. Fadok, V. A., Voelker, D. R., Campbell, P. A., Cohen, J. J., Bratton, D. L., Henson, P. M. (1992) J. Immunol. 148, 2207-2216[Abstract/Free Full Text]
  5. Bratton, D. L., Fadok, V. A., Richter, D. A., Kailey, J. M., Guthrie, L. A., Henson, P. M. (1997) J. Biol. Chem. 272, 26159-26165[Abstract/Free Full Text]
  6. Hübscher, H. G., Dils, R. R., and Pover, W. F. R. (1959) Biochim. Biophys. Acta 36, 518-525[Medline] [Order article via Infotrieve]
  7. Okada, M., Matsuzaki, H., Shibuya, I., and Matsumoto, K. (1994) J. Bacteriol. 176, 7456-7461[Abstract]
  8. Letts, V. A., Klig, L. S., Bae-Lee, M., Carman, G. M., Henry, S. A. (1983) Proc. Natl. Acad. Sci. U. S. A. 80, 7279-7283[Abstract]
  9. Nikawa, J., Tsukagoshi, Y., Kodaki, T., and Yamashita, S. (1987) Eur. J. Biochem. 167, 7-12[Abstract]
  10. Kuge, O., Nishijima, M., and Akamatsu, Y. (1991) J. Biol. Chem. 266, 24184-24189[Abstract/Free Full Text]
  11. Saito, K., Kuge, O., Akamatsu, Y., and Nishijima, M. (1997) FEBS Lett. 395, 262-266[CrossRef]
  12. Vance, J. E. (1990) J. Biol. Chem. 265, 7248-7256[Abstract/Free Full Text]
  13. Kuge, O., Saito, K., and Nishijima, M. (1997) J. Biol. Chem. 272, 19133-19139[Abstract/Free Full Text]
  14. Voelker, D. R., and Frazier, J. L. (1986) J. Biol. Chem. 261, 1002-1008[Abstract/Free Full Text]
  15. Dennis, E. A., and Kennedy, E. P. (1972) J. Lipid Res. 13, 263-267[Abstract/Free Full Text]
  16. Shiao, Y.-J., Lupo, G., and Vance, J. E. (1995) J. Biol. Chem. 270, 11190-11198[Abstract/Free Full Text]
  17. Kennedy, E. P., and Weiss, S. B. (1956) J. Biol. Chem. 222, 193-214[Free Full Text]
  18. Sundler, R. (1973) Biochim. Biophys. Acta 306, 218-226[Medline] [Order article via Infotrieve]
  19. Tijburg, L. B. M., Geelen, M. J. H., van Golde, L. M. G. (1989) Biochem. Biophys. Res. Commun. 160, 1275-1280[Medline] [Order article via Infotrieve]
  20. Zelinski, T. A., and Choy, P. C. (1982) Can. J. Biochem. 60, 817-823[Medline] [Order article via Infotrieve]
  21. Yeung, S. K. F., and Kuksis, A. (1976) Lipids 11, 498-505[Medline] [Order article via Infotrieve]
  22. Voelker, D. R. (1984) Proc. Natl. Acad. Sci. U. S. A. 81, 2669-2673[Abstract]
  23. Miller, M. A., and Kent, C. (1986) J. Biol. Chem. 261, 9753-9761[Abstract/Free Full Text]
  24. Kano-Sueoka, T., Cohen, D. M., Yamaizumi, Z., Nishimura, S., Mori, M., and Fujiki, H. (1979) Proc. Natl. Acad. Sci. U. S. A. 76, 5741-5744[Abstract]
  25. Murakami, H., Masui, H., Sato, G. H., Sueoka, N., Chow, T. P., Kano-Sueoka, T. (1982) Proc. Natl. Acad. Sci. U. S. A. 79, 1158-1162[Abstract]
  26. Kano-Sueoka, T., Errick, J. E., King, D., and Walsh, L. A. (1983) J. Cell. Physiol. 117, 109-115[Medline] [Order article via Infotrieve]
  27. Sasaki, H., Kume, H., Nemoto, A., Narisawa, S., and Takahashi, N. (1997) Proc. Natl. Acad. Sci. U. S. A. 94, 7320-7325[Abstract/Free Full Text]
  28. Menon, A. K., and Stevens, V. L. (1992) J. Biol. Chem. 267, 15277-15280[Abstract/Free Full Text]
  29. Sambrook, J., Fritsch, E. F., and Maniatis, T. (1989) Molecular Cloning: A Laboratory Manual, 2nd. Ed., pp. 2.72-2.81, Cold Spring Harbor Laboratory, Cold Spring Harbor, NY
  30. Sanger, F., Nicklen, S., and Coulson, A. R. (1977) Proc. Natl. Acad. Sci. U. S. A. 74, 5462-5467
  31. Yao, Z., and Vance, D. E. (1988) J. Biol. Chem. 263, 2998-3004[Abstract/Free Full Text]
  32. Chen, C., and Okayama, H. (1987) Mol. Biol. Cell 7, 2745-52
  33. Vance, J. E., and Vance, D. E. (1988) J. Biol. Chem. 263, 5898-5909[Abstract/Free Full Text]
  34. Porter, T. J., and Kent, C. (1992) Methods Enzymol. 209, 134-146[Medline] [Order article via Infotrieve]
  35. Tijburg, L. B. M., Vermeulen, P. S., van Golde, L. M. B. (1992) Methods Enzymol. 209, 258-266[Medline] [Order article via Infotrieve]
  36. Bagu, J. R., Sonnischen, F. D., Williams, D., Anderson, R. J., Sykes, B. D., Holmes, C. F. B. (1995) Nat. Struct. Biol. 2, 114-116[Medline] [Order article via Infotrieve]
  37. Bligh, E. G., and Dyer, W. J. (1959) Can. J. Biochem. Physiol. 37, 911-917
  38. Zhou, X., and Arthur, G. (1992) J. Lipid Res. 33, 1233-1236[Abstract]
  39. Macala, L. J., Ku, R. K., and Ando, S. (1983) J. Lipid Res. 24, 1243-1250[Abstract]
  40. Kyte, J., and Doolittle, R. F. (1982) J. Mol. Biol. 157, 105-132[Medline] [Order article via Infotrieve]
  41. Schutze, M.-P., Peterson, P. A., and Jackson, M. R. (1994) EMBO J. 13, 1696-1705[Abstract]
  42. Nikawa, J.-I., Tsukagoshi, Y., Kodaki, T., and Yamashita, S. (1987) Eur. J. Biochem. 167, 7-12[Abstract]
  43. Raetz, C. R. H., Larson, T. J., and Dowhan, W. (1977) Proc. Natl. Acad. Sci. U. S. A. 74, 1412-1416[Abstract]
  44. Tijburg, L. B. M., Geelen, M. J. H., van Golde, L. M. G. (1989) Biochim. Biophys. Acta 1004, 1-19[Medline] [Order article via Infotrieve]
  45. Tijburg, L. B. M., Houweling, M., Geelen, M. J. H., van Golde, L. M. G. (1987) Biochim. Biophys. Acta 922, 184-190[Medline] [Order article via Infotrieve]
  46. Sundler, R., and Akesson, B. (1975) J. Biol. Chem. 250, 3359-3367[Abstract]
  47. Tijburg, L. B. M., Schuurmans, E. A. J. M., Geelen, M. J. H., van Golde, L. M. G. (1987) Biochim. Biophys. Acta 919, 49-57[Medline] [Order article via Infotrieve]
  48. Tijburg, L. B. M., Houweling, M., Geelen, M. J. H., van Golde, L. M. G. (1989) Biochem. J. 257, 645-650[Medline] [Order article via Infotrieve]
  49. Walkey, C. J., Kalmar, G. B., and Cornell, R. B. (1994) J. Biol. Chem. 269, 5742-5749[Abstract/Free Full Text]
  50. Ohta, A., Waggoner, K., Louie, K., and Dowhan, W. (1981) J. Biol. Chem. 256, 2219-2225[Free Full Text]
  51. Voelker, D. R. (1989) J. Biol. Chem. 264, 8019-8025[Abstract/Free Full Text]
  52. Samborski, R. W., and Vance, D. E. (1993) Biochim. Biophys. Acta 1167, 15-21[Medline] [Order article via Infotrieve]
  53. Vermeulen, P. S., Tijburg, L. B. M., Geelen, M. J. H., van Golde, L. M. G. (1993) J. Biol. Chem. 268, 7458-7464[Abstract/Free Full Text]
  54. Schneider, W., Fiscus, W. G., and Lawler, J. A. (1986) Anal. Biochem. 14, 121-134
  55. van Hellemond, J. J., Slot, J. W., Geelen, M. J., van Golde, L. M., Vermeulen, P. S. (1994) J. Biol. Chem. 269, 15415-15418[Abstract/Free Full Text]
  56. Nakashima, A., Hosaka, K., and Nikawa, J. (1997) J. Biol. Chem. 272, 9567-9572[Abstract/Free Full Text]
  57. Kalmar, G. B., Kay, R. J., Lachance, A., Aebersold, R., and Cornell, R. B. (1990) Proc. Natl. Acad. Sci. U. S. A. 87, 6029-6033[Abstract]
  58. Craig, L., Johnson, J. E., and Cornell, R. B. (1994) J. Biol. Chem. 269, 3311-3317[Abstract/Free Full Text]
  59. Lim, P., Cornell, R., and Vance, D. E. (1986) Biochem. Cell Biol. 64, 692-698[Medline] [Order article via Infotrieve]
  60. Araki, W., and Wurtman, R. J. (1997) Proc. Natl. Acad. Sci. U. S. A. 94, 11946-11950[Abstract/Free Full Text]
  61. Jamil, H., Utal, A. K., and Vance, D. E. (1992) J. Biol. Chem. 267, 1752-1760[Abstract/Free Full Text]


Copyright © 1998 by The American Society for Biochemistry and Molecular Biology, Inc.