©1995 by The American Society for Biochemistry and Molecular Biology, Inc.
Phosphatidylcholine Turnover in Activated Human Neutrophils
AGONIST-INDUCED CYTIDYLYLTRANSFERASE TRANSLOCATION IS SUBSEQUENT TO PHOSPHOLIPASE D ACTIVATION (*)

Hélène Tronchère (§) , Valérie Planat , Michel Record (¶) , Franois Tercé , Gérard Ribbes , Hugues Chap

From the (1) Institut National de la Santé et de la Recherche Médicale, Unité 326, Phospholipides Membranaires, Signalisation Cellulaire et Lipoprotéines, Hôpital Purpan, 31059 Toulouse Cedex, France

ABSTRACT
INTRODUCTION
EXPERIMENTAL PROCEDURES
RESULTS
DISCUSSION
FOOTNOTES
REFERENCES

ABSTRACT

Phosphatidylcholine synthesis and degradation are tightly regulated to assure a constant amount of the phospholipid in cellular membranes. The chemotactic peptide fMLP and the phorbol ester, phorbol 12-myristate 13-acetate, are known to stimulate phosphatidylcholine degradation by phospholipase D in human neutrophils. fMLP alone triggered phosphatidylcholine breakdown into phosphatidic acid, but did not stimulate phosphatidylcholine synthesis or activation of the rate-limiting enzyme CTP:phosphocholine cytidylyltransferase. Adding cytochalasin B to fMLP led to some conversion of phosphatidic acid into diglyceride, and fMLP was then able to trigger choline incorporation into phosphatidylcholine, and cytidylyltransferase translocation from cytosol to membranes. Inhibition of phosphatidylcholine-phospholipase D activation with tyrphostin led to inhibition of choline incorporation. Therefore, phosphatidic acid-derived diglyceride but not phosphatidic acid alone was effective to promote cytidylyltransferase translocation. With phorbol 12-myristate 13-acetate as agonist, and by selective labeling of phosphatidylinositol and phosphatidylcholine, we demonstrated that only phosphatidylcholine-derived diglyceride participated in cytidylyltransferase translocation. Oleic acid stimulated phosphatidylcholine synthesis, but induced a weak increase in diglyceride and a slight cytidylyltransferase translocation, and did not stimulate phospholipase D activity. Our data established that only diglyceride derived from phosphatidylcholine degradation by the phospholipase D/phosphatidate phosphatase pathway are required for agonist-induced cytidylyltransferase translocation and subsequent choline incorporation into phosphatidylcholine.


INTRODUCTION

The presence of a phosphatidylcholine cycle related to the sn-2 position of the glycerol backbone through the deacylation/reacylation pathway has been characterized in the human neutrophil (1) . Evidence for cycling degradation/resynthesis reactions concerning the sn-3 position have also been reported in the work of Daniel et al.(2) and proposed by Pelech and Vance (3) as a basal turnover pathway, but were rarely investigated upon cell activation. The CDP-choline pathway accounts for the main production of phosphatidylcholine in mammalian cells: phosphatidylcholine is synthesized from external choline in three enzymatic steps involving, respectively, choline kinase (EC 2.7.1.32), CTP:phosphocholine cytidylyltransferase (EC 2.7.7.15), and choline phosphotransferase (EC 2.7.8.2). Cytidylyltransferase is the key regulatory enzyme of the pathway and its activity can be regulated by different factors (4) . Two forms of the enzyme are present in cells: a non-active cytosolic form and a highly active membrane-bound form (5) . Diglyceride, formed in cells challenged with PMA,() has been pointed out to control cytidylyltransferase translocation (6) . Whether phosphatidylcholine degradation into diglyceride occurs prior or subsequent to enhanced [H]choline incorporation via the CDP-choline pathway has been a matter of debate (6, 7) .

The present investigation has been undertaken on human neutrophils whose phosphatidylcholine degradation at the sn-3 position by phospholipase D has been characterized (8, 9, 10, 11) . Earlier studies on phosphatidylcholine synthesis in this cell model were mainly related to methylation of phosphatidylethanolamine (12, 13) , to the effect of anti-tubulin or anti-microfilament agents (14, 15) , or to phosphatidylcholine turnover during phagocytosis (16) . In the latter study, activation of choline phosphotransferase was reported. However, cytidylyltransferase has not been studied yet in a hematopoietic cell line. This study has been conducted using three different compounds: oleic acid, the phorbol ester PMA, and the receptor-dependent chemotactic peptide, fMLP. Oleic acid has been shown to be a potent activator of cytidylyltransferase translocation and has been studied as such in various models (17, 18) . PMA stimulates phosphatidylcholine turnover in several cell types (2, 19, 20) . Stimulation of phosphatidylcholine synthesis appears to involve protein kinase C (21) , without implicating a direct regulation of cytidylyltransferase by this kinase (6, 22) . Moreover, PMA stimulates phosphatidylcholine degradation via phospholipase D, whose activation partly involves protein kinase C (23) . fMLP binds to a specific seven-pass transmembrane receptor on human neutrophils and has been shown to induce phosphatidylcholine degradation through phospholipase D activation (11) , since neutrophils are lacking phosphatidylcholine-specific phospholipase C (24) . Although the fMLP receptor is coupled to G-proteins, phospholipase D activation is dependent upon the activity of tyrosine kinases (25) .

We now report that stimulation of [H]choline incorporation into phosphatidylcholine, triggered by PMA and fMLP, depends only on the presence of diglyceride formed from phosphatidylcholine-derived phosphatidic acid. Our data provide evidence for an ``activated'' phosphatidylcholine cycle in human neutrophils triggered by receptor and non-receptor agonists, and indicates a coupling between phospholipase D and cytidylyltransferase activation. Such a coupling is not required for oleic acid which triggers [H]choline incorporation without phospholipase D activation.


EXPERIMENTAL PROCEDURES

Chemicals and Products

[methyl-H]Choline chloride (2.89 TBq (78Ci)/mmol), phospho[methyl-C]choline-ammonium salt (2.22 GBq (60 mCi)/mmol), [-P]ATP (110 TBq (3000 Ci)/mmol), [5,6,8,9,11,12,14,15-H]arachidonic acid (7.73 TBq (209Ci)/mmol), and 1-O-[H]alkyl-2-lyso-sn-glycero-3-phosphocholine ([H]alkyl-labeled lyso-PAF) (80 Ci/mmol) were purchased from the Radiochemical Center (Amersham, Bucks, United Kingdom). The [H]choline-labeled lyso-PAF, i.e. 1-O-hexadecyl-2-lyso-sn-glycero-3-phospho[N-methyl-H]choline (3.07 TBq (83 Ci)/mmol) was specially prepared by Amersham (UK). Percoll was obtained from Pharmacia (Uppsala, Sweden). PMA, 4-O-methyl-PMA, fMLP, cytochalasin B, tyrphostin 47, oleic acid, digitonin, and all other chemicals were from Sigma. All stock solutions were prepared in dimethyl sulfoxide whose final concentrations in cell incubation and assays never exceeded 0.02% (v/v). When oleic acid was dissolved in ethanol, final ethanol concentration was 1% (v/v).

Composition of Buffers

Buffer A contained 20 mM Hepes, pH 7.4, 137 mM NaCl, 2.6 mM KCl, 5.5 mM glucose, 10 µM choline; and buffer B contained buffer A without choline. The digitonin buffer consisted of 0.2 mg/ml digitonin, 10 mM Tris-HCl, pH 7.4, 0.25 M sucrose, 0.5 mM phenylmethylsulfonyl fluoride.

Neutrophil Isolation

Buffy coats were obtained from healthy donors of the local blood bank (Centre Régional de Transfusion Sanguine, Toulouse, France). Human neutrophils were separated by using a slight modification of the method previously described (26) . Briefly, a leukocyte-rich supernatant was obtained following addition of a 2% (w/v) dextran solution in isotonic NaCl to a buffy coat. After centrifugation, remaining erythrocyte contamination of the leukocyte pellet was eliminated with suspension for 10 min in cold isotonic NHCl. Separation of leukocytes across a Percoll gradient gave a final cell population of more than 98% neutrophils, as assessed with May-Grünswald-Giemsa staining.

Cell Stimulation

In experiments dealing with choline labeling, 10 cells were incubated in buffer A with 1 µCi of [H]choline (final specific radioactivity of 0.4 µCi/nmol) for 20 min prior addition of stimuli (oleic acid, PMA, or 4-O-methyl-PMA, fMLP). At each incubation time, 0.5 ml of cell suspension (2 10 cells) was harvested and pelleted (2,800 g for 1 min) using an MSE microcentrifuge (Kontron Instruments). The cell pellet was extracted according to Bligh and Dyer (27) and radioactivity of the organic phase was determined. This radioactivity corresponded to phosphatidylcholine labeling only, as checked by analysis on thin layer chromatography (TLC). In some experiments, lactate dehydrogenase activity was measured in the supernatant of incubation.

When neutrophils were labeled either with [H]arachidonic acid or [H]lyso-PAF, 10 cells were incubated for 30 min with 1 µCi of the tritiated compound, washed with buffer A containing 2.5% (w/v) bovine serum albumin, and finally resuspended in buffer A.

Pulse-Chase Experiments

Neutrophils were prelabeled in buffer A containing [H]choline (1 µCi/1 10 cells) for 30 min. After removal of the labeling medium, cells were rinsed twice and incubated in medium without labeled precursor and in the presence of the agonists. At each incubation time, 0.5 ml of cell suspension was harvested, extracted according to Bligh and Dyer (27) , and radioactivity from the organic phase was determined. The labeled aqueous choline metabolites were separated by TLC in methanol, 0.6% NaCl, NHOH (50/50/5, v/v) (28) . The spots were identified, scraped off, and counted for radioactivity.

Digitonin Permeabilization of Cells

Digitonin permeabilization was performed as described by Pelech et al. (29). Briefly, neutrophils were rinsed twice with cold incubation buffer B after treatment with the agonists and permeabilized with digitonin buffer at 4 °C for the indicated times. After centrifugation at 13,000 g for 1 min, the supernatant (released cytosolic content) was kept at 4 °C before cytidylyltransferase assay. Meanwhile the pellet (containing cell ghosts) was resuspended in 10 mM Tris-HCl, pH 7.4, 0.25 M sucrose, 0.5 mM phenylmethylsulfonyl fluoride, and sonicated with three bursts of 1 s at 30% power output with a microtip probe equipped-sonicator (Heat Systems, Ultrasonics Inc., model W-225 R).

Cytidylyltransferase Assay

CTP:phosphocholine cytidylyltransferase activity was assayed as described previously (30) . The incubation mixture contained 20 mM Tris succinate, pH 7.8, 6 mM MgCl, 8 mM CTP, 4 mM phospho-[methyl-C]choline (0.5 mCi/mmol), and up to 300 µg of protein from cytosol or membrane fractions. A sonicated suspension of total lipid extract (1 mM lipid phosphorus) from Krebs-II cells was added to assay the cytosolic enzyme. The incubation was carried out for 30 min at 37 °C and stopped by boiling in the presence of unlabeled phosphocholine (200 mM final concentration). [C]CDP-choline was separated and measured as described (28).

Mass Measurement of Diglyceride

Neutrophils were incubated in buffer B for the indicated times in the presence of agonists and then extracted according to the method of Bligh and Dyer (27) . An aliquot of the lipid fraction was dried under nitrogen and resolubilized with 20 µl of 7.5% (w/v) octyl--D-glucoside, 5 mM cardiolipin, 1 mM diethylenetriaminepentaacetic acid. Diglyceride amounts were determined as described by Wright et al.(31) , a modification of the method of Preiss et al.(32) , using Escherichia coli diglyceride kinase (Lipidex, Inc.) and [-P]ATP.

In Situ Determination of Phospholipase D Activity

To analyze the generation of labeled phosphatidic acid and diglyceride, cells were labeled with [H]alkyl-lyso-GPC (1 µCi/10 cells) for 30 min at 37 °C (9, 11) , then stimulated with the appropriate compound. Lipids were extracted using Bligh and Dyer procedure (27) , and phosphatidylcholine breakdown products were separated on silica plates according to Olson et al.(33) .

Phospholipase D activity was also monitored with the choline-labeled lyso-PAF, which was reacylated in cells into alkylacyl-glycerophospho[H]choline. Activation of the phospholipase was represented by the release of free [H]choline in the upper phase of the Bligh and Dyer extract. In that case, 10 cells were labeled with 1 µCi of the radioactive precursor for 30 min prior to stimulation. Inhibition of phospholipase D activation by tyrphostin was obtained in cells preincubated for 5 min with a final concentration of 100 µM of the compound before stimulation.

Miscellaneous Determinations

Protein was determined according to the method of Lowry et al.(34) in the presence of sodium dodecyl sulfate (0.07%, w/v), using bovine serum albumin as a standard. Lactate dehydrogenase activity and N-acetyl--D-glucosaminidase activity were measured as described previously (35, 36) .

Data Presentation

Results are expressed as the average ± S.E. of three separate experiments, or the average of two experiments with less than 10% variance.


RESULTS

Effect of Oleic Acid and Phorbol Ester on [H]Choline Incorporation into Phosphatidylcholine (Fig. 1)-Human neutrophils were incubated with [H]choline and various concentrations of oleic acid or PMA. Results reported in Fig. 1A indicate a sharp dose dependence of stimulated [H]choline incorporation when exposed to the fatty acid. A 5-fold stimulation was obtained at concentrations of oleic acid between 40 and 60 µM, within the range of non-lytic concentrations. The dose dependence was very similar to our previous results on HL-60 cells (37) .


Figure 1: Concentration-dependent incorporation of [H]choline into phosphatidylcholine in the presence of oleic acid (A) or PMA (B). Neutrophils were incubated with [H]choline for 30 min in the presence of oleic acid (Panel A, ]), or for 20 min in the presence of PMA (Panel B, ) or 4-O-methyl-PMA (Panel B, ). Radioactivity incorporated into phosphatidylcholine was measured as described under ``Experimental Procedures.'' Cell integrity was monitored with release of lactate dehydrogenase (). Results are expressed as average ±S.E. of three determinations (A), or average of two determinations (B).



As shown in Fig. 1B, PMA stimulated [H]choline incorporation by 2.5-fold with an optimal effect between 100 and 500 nM, a concentration range used in many studies. On the other hand, 4-O-methyl-PMA, a phorbol ester which does not activate protein kinase C (38) , did not produce any change in phosphatidylcholine labeling.

Cytidylyltransferase Catalyzes the Rate-limiting Step of the CDP-choline Pathway in Human Neutrophils (Fig. 2)

We next investigated the labeling of the different choline derivatives in cells challenged with oleic acid or PMA. As compared to untreated cells, pulse-chase experiments performed with oleic acid revealed a strong decrease in phosphocholine labeling, and an increase in CDP-choline labeling (Fig. 2A). Free internalized [H]choline also showed a time-dependent decrease but no change between control and treated cells was observed. Labeling of phosphatidylcholine was clearly increased in the presence of 40 µM oleic acid. The variations observed for phosphocholine radioactivity were opposite to those for CDP-choline and phosphatidylcholine. This is consistent with cytidylyltransferase being the rate-limiting enzyme of the CDP-choline pathway in our cell model (30) .


Figure 2: Pulse-chase experiments with [H]choline in the presence of oleic acid (A) or PMA (B). Neutrophils were prelabeled for 30 min with [H]choline in Buffer A, separated in two pools, rinsed twice, and incubated in the absence () or presence of 40 µM oleate (Panel A, ) or 500 nM PMA (Panel B, ). At the indicated times, cells were harvested and lipids extracted. Radioactivity of aqueous choline metabolites and phosphatidylcholine was determined: (a) free internalized choline; (b) phosphocholine; (c) CDP-choline; and (d) phosphatidylcholine. Results are expressed as average of two (A) or three determinations (B) ±S.E.



A similar pattern was also observed using PMA as an agonist (Fig. 2B). [H]Choline levels remained constant for 30 min with no change between control and treated cells. Phosphocholine labeling decreased after a lag period of 5 min. A similar lag time was noticed for the enhancement of phosphatidylcholine radioactivity, whereas no clear-cut modification of CDP-choline levels was evident. This might indicate a concomitant stimulation of choline phosphotransferase, as we previously reported for phospholipase C-treated Krebs-II cells (30) . Similar results to those reported with PMA were obtained with fMLP as an agonist (not shown).

Comparative Effect of Oleic Acid and Agonists on Phosphatidylcholine Synthesis and Cytidylyltransferase Translocation ( Fig. 3 and Fig. 4)

We have performed a time course study of [H]choline incorporation in the presence of the chemotactic peptide fMLP, PMA, or 4-O-methyl-PMA. The stimulation of [H]choline incorporation into phosphatidylcholine was better observed when cells were preincubated for 20 min with [H]choline prior to the addition of cytochalasin B and fMLP. Neither cytochalasin B nor fMLP alone had any stimulating effect (Fig. 3A). The ratio of [H]choline incorporation in cells stimulated with fMLP plus cytochalasin B versus control was maximal at 5 min. The phorbol ester PMA was also clearly able to stimulate [H]choline incorporation into phosphatidylcholine, after a lag time of 5 min (Fig. 3B). The analog 4-O-methyl-PMA showed no effect. The ratio of choline incorporation in cells stimulated with PMA versus control continuously increased during the 20-min time course.


Figure 3: Time course of [H]choline incorporation into phosphatidylcholine in neutrophils stimulated with fMLP and PMA. Neutrophils were preincubated with [H]choline for 20 min, then incubated in the absence () or presence of: Panel A, 1 µM fMLP (), 5 µM cytochalasin B (), or 1 µM fMLP + 5 µM cytochalasin B (); Panel B, 500 nM PMA () or 500 nM 4-O-methyl-PMA (). Radioactivity in the organic phase was determined, and results were expressed as average of two determinations.




Figure 4: Cytidylyltransferase redistribution upon cell activation. Cell activation was performed for 20 min with optimum concentration of agonists, and in the presence of cytochalasin B for fMLP. Cytidylyltransferase activity was measured in the digitonin-release medium (cytosolic form of the enzyme) and in the presence of exogenous lipids (Panel A). The activity remaining in the cell ghosts was determined in the absence of added lipids, and accounted for the membranous form of the enzyme (Panel B). The sum of cytosolic plus particulate activities is represented in Panel C. Results are expressed as picomolemin and are mean ± S.E. of three different experiments. Cont: control (untreated) cells.



The relative distribution of cytidylyltransferase between cytosol and membranes was investigated after 20 min incubation and using two experimental approaches giving similar results: cell disruption with nitrogen cavitation followed with centrifugation to separate particulate and cytosolic fractions (not shown), or cell permeabilization with digitonin (Fig. 4). Optimal digitonin concentration and permeabilization time were determined (data not shown) by measuring release of lactate dehydrogenase activity from cytosol and by measuring N-acetyl--D-glucosaminidase activity as a marker of granule integrity (see ``Experimental Procedures''). Results reported in Fig. 4display the amount of cytidylyltransferase retained in the cell (particulate form) or released into supernatant (cytosolic form).

In the presence of PMA, a lower amount of cytidylyltransferase was recovered in the cytosolic fraction of permeabilized cells, as compared to controls. (Fig. 4A). By contrast the activity in the particulate was increased by 16-fold (Fig. 4B). However, the sum of total activity in particulate plus cytosol was higher than the value in non-treated cells (Fig. 4C), indicating that binding of cytidylyltransferase on membranes leads to an overactivation of the enzyme, about 50% above the basal value. Nevertheless, PMA induced the translocation to membranes of a high amount of cytidylyltransferase. Oleic acid treatment had only a slight effect on cytidylyltransferase translocation. When the cells were stimulated with fMLP plus cytochalasin B, we observed the binding of cytidylyltransferase to membranes (Fig. 4B), and no variation in the sum of total activity between cytosol and particulate as compared to control cells (Fig. 4C). Thus, our data show a modification of cytidylyltransferase distribution between cytosol and membranes induced with PMA and fMLP, in contrast to oleic acid treatment that induced a weak redistribution of the enzyme. Relationship between Phosphatidylcholine Breakdown and Resynthesis (Fig. 5-8)-To investigate the mechanisms involved in the regulation of cytidylyltransferase translocation, we have analyzed the amount of diglyceride produced in the different experimental conditions (Fig. 5). Basal amounts of diglyceride varied from about 100 to 300 pmol/10 cells, depending upon the batch of neutrophils; such variations being in the range of literature data. Oleic acid induced only a weak formation of diglyceride (Fig. 5A). In contrast, diglyceride increased continuously after a 5-min lag time in PMA-treated neutrophils, whereas 4-O-methyl-PMA had no effect (Fig. 5B). A rapid 2-fold increase in diglyceride mass was only noticed with fMLP plus cytochalasin B, reaching a maximum at 5 min and decreasing slowly thereafter (Fig. 5C). In this case, the maximum of diglyceride formation corresponded to the maximum enhancement of [H]choline incorporation, as determined by the ratios between stimulated and resting cells for diglyceride and phosphatidylcholine labeling (not shown). Also, the kinetics of PMA-induced diglyceride formation (Fig. 5B) paralleled that of [H]choline incorporation in cells stimulated with the phorbol ester (Fig. 3B).


Figure 5: Time course of diglyceride formation in activated neutrophils. Neutrophils were incubated for the indicated times in the absence () or presence of: Panel A, 40 µM oleic acid (); Panel B, 500 nM PMA () or 500 nM 4-O-methyl-PMA (); Panel C, 1 µM fMLP (), 5 µM cytochalasin B (), or 1 µM fMLP + 5 µM cytochalasin B (). Following lipid extraction, diglyceride amount was measured as described under ``Experimental Procedures.'' Results are expressed as average of two determinations.



Because phosphatidylcholine degradation was well documented in human neutrophils (8, 9, 10, 11) , results from Fig. 5 prompted us to investigate a possible relationship between phosphatidylcholine degradation and synthesis through the generation of diglyceride (9, 10) . In Fig. 6A, stimulation of neutrophils with fMLP led only to the formation of phosphatidylcholine-derived phosphatidic acid. In contrast, simultaneous addition of fMLP and cytochalasin B enhanced phosphatidic acid generation, and triggered the formation of phosphatidic acid-derived diglyceride (Fig. 6B). Cytochalasin B also increased the release of water-soluble radioactivity from endogenous alkylacyl-glycerophospho[H]choline (Fig. 6C). When passed through an anion exchange column (Dowex 50), the radioactivity was found to correspond to [H]choline only (not shown), which is in agreement with the absence of a phospholipase C acting on phosphatidylcholine in the human neutrophil (24) . Therefore addition of the priming compound cytochalasin B to the fMLP agonist triggered both [H]choline release and phosphatidylcholine-derived diglyceride production, indicating an activation of the phospholipase D/phosphatidic acid phosphatase pathway.


Figure 6: Kinetics of phospholipase D-mediated phosphatidylcholine breakdown products from endogenously labeled [H]alkylacyl-GPC or alkylacylglycerophospho[H]choline. Cells were prelabeled with [H]alkyl-lyso-glycerophosphocholine (Panels A, B, D, E, and F) or alkyl-lyso-glycerophospho[H]choline (Panel C). Generation of [H]alkyl-labeled phosphatidic acid () and diglyceride () was monitored in cells stimulated with: Panel A, 1 µM fMLP; Panel B, 1 µM fMLP + 5 µM cytochalasin B; Panel D, 100 nM PMA; Panel E, 40 µM oleic acid. When oleic acid (40 µM) was added in the presence of ethanol (1% final v/v) no alkyl-labeled phosphatidylethanol (Panel F) was recovered (), as compared to cells incubated with ethanol only (). Results are expressed as percent of total radioactivity from 10 cells (A, B, D-F) and are average of two determinations. Panel C, [H]choline release from alkylacyl-glycerophospho[H]choline was checked in the presence of fMLP () or fMLP + cytochalasin B (), in the conditions of Panel B. Results are expressed as disintegrations/min recovered in the water phase of lipid extract from 10 labeled cells. Average of two determinations.



When neutrophils were stimulated with the non-receptor agonist PMA, both phosphatidic acid and diglyceride were produced (Fig. 6D). Phosphatidic acid formation preceded that of diglyceride whose onset time was between 2.5 and 5 min, a time comparable to that required for PMA-induced [H]choline incorporation (Fig. 3B). The non-agonist compound, oleic acid, was not an activator of phospholipase D, as monitored with the lack of phosphatidic acid and phosphatidylethanol formation (Fig. 6, E and F). As a consequence, no phosphatidic acid-derived diglyceride was produced, which appeared consistent with the weak changes in the mass amount of total diglyceride (Fig. 5A) and the slight cytidylyltransferase translocation (Fig. 4).

To better assess whether phosphatidylcholine-derived diglyceride alone controlled cytidylyltransferase translocation, we differentially labeled phosphatidylinositol and phosphatidylcholine, and followed their respective conversion into diglyceride upon cell stimulation with PMA (Fig. 7). Cell labeling with [H]arachidonic acid was distributed mainly into phosphatidylinositol as reported (16) . In contrast, 90% of [H]lyso-PAF was incorporated into phosphatidylcholine as we previously observed (9, 11) . Addition of PMA induced no variation in the level of arachidonyl-labeled diglyceride (Fig. 7A), at variance with the results observed for [H]alkyl-labeled diglyceride, whose kinetics of labeling was similar to that of total diglyceride (Fig. 5B). Comparison between Fig. 7, B and C, showed that the association of cytidylyltransferase with the membrane paralleled the phosphatidylcholine-derived diglyceride formation.


Figure 7: Comparison between time course generation of [H]arachidonyl or [H]alkyl-labeled diglyceride and membrane association of cytidylyltransferase. Cells were labeled with either [H]arachidonic acid or [H]alkyl-lyso-GPC, and stimulated with 100 nM PMA. Panel A, [H]arachidonyl-labeled diglyceride (); Panel B, [H]alkyl-labeled diglyceride () and control cells (). Results are expressed as percent of total radioactivity and are average of two determinations. Panel C, specific activity of membrane-bound cytidylyltransferase in PMA-treated cells () as compared to non-treated cells (). Average of two determinations.



With the receptor agonist fMLP, phosphatidylinositol-derived diglyceride would also have been produced since the agonist also stimulates the phosphatidylinositol-phospholipase C (39) . To monitor the incidence of phospholipase D-mediated phosphatidylcholine hydrolysis on the phospholipid resynthesis, we blocked phospholipase D activation with a tyrosine kinase inhibitor, tyrphostin. This compound lead to a strong inhibition of the [H]choline release (Fig. 8A). The effect of fMLP plus cytochalasin B on stimulation of [H]choline incorporation was monitored in a parallel series of samples. In control cells, time course of [H]choline incorporation was slower than the phospholipase D-mediated release of the base, since maximum incorporation occurred at 15 min (Fig. 8B), whereas maximum release was noticed at 1 min (Fig. 8A). As a consequence of phospholipase D inhibition, [H]choline incorporation in fMLP-stimulated cells no longer occurred (Fig. 8B). Therefore, phospholipase D-mediated choline release from phosphatidylcholine was required prior to [H]choline incorporation into this phospholipid when neutrophils were activated with fMLP. The choline released upon cell activation almost leveled off from 1 to 25 min (Fig. 8A), suggesting that this pool of free choline was not reutilized for the phosphatidylcholine resynthesis monitored in Fig. 8B.


Figure 8: Comparative effect of a tyrosine kinase inhibitor on phosphatidylcholine breakdown and resynthesis upon cell stimulation with fMLP. Panel A, release of [H]choline from endogenous alkylacyl-glycerophospho[H]choline in cells incubated with 1 µM fMLP + cytochalasin B () or 1 µM fMLP + cytochalasin B following a 5-min treatment with 100 µM tyrphostin (), as compared to control cells (). Data represent the radioactivity recovered in the aqueous phase. Panel B, incorporation of free labeled choline into phosphatidylcholine from another set of cells, in the conditions of Panel A: fMLP-activated cells (), or fMLP-activated cells + tyrphostin (), as compared to control cells (). Data represent the radioactivity recovered in the organic phase. Results are average of two determinations.




DISCUSSION

In the present work we have investigated activation of a phosphatidylcholine cycle triggered with agonists and oleic acid in a terminally differentiated cell. Hydrolysis of phosphatidylcholine, induced with a variety of agonists, has been well characterized in this cell type (8, 9, 10, 11) . Phosphatidylcholine has been identified as an alternative source for phosphatidic acid and diglyceride as second messengers (5, 8, 9, 10, 11) , and an activated phosphatidylcholine cycle, analogous to that of phosphoinositide, appears necessary to reconstitute the initial amount of cell phosphatidylcholine (3) . However, resynthesis of phosphatidylcholine from diglyceride is regulated by the amount of CDP-choline, which is produced by the enzyme cytidylyltransferase. Therefore, the present study has been focused on cytidylyltransferase activation and its possible relationship to phosphatidylcholine degradation.

We show that oleic acid and PMA stimulate [H]choline incorporation into phosphatidylcholine (Fig. 1). Moreover, by performing pulse-chase experiments, we demonstrate that cytidylyltransferase is the rate-limiting enzyme for the CDP-choline pathway for phosphatidylcholine synthesis in human neutrophils (Fig. 2). Two different patterns of CDP-choline labeling are observed in Fig. 2and are consistent with previous observations on growing cells (30, 40) . So far, the CDP-choline pathway has not been carefully studied in terminally differentiated cells, and specifically in hematopoietic ones.

Preincubation of the cells with the labeled choline prior to addition of the agonist is required to observe a net effect of fMLP as well as PMA. It suggests that a specific pool of choline might be involved, or that some channeling would be required (41) . The increase of phosphatidylcholine radioactivity in fMLP-stimulated neutrophils was strictly dependent on the presence of cytochalasin B (Fig. 3A). Cytochalasin B has been widely used in neutrophil studies, since it enhances cellular responses such as degranulation or oxidative burst (42) . In the presence of cytochalasin B, the fMLP-induced phosphatidylcholine synthesis appears to level off rapidly, whereas with PMA, enhanced [H]choline incorporation appears to be a long lasting event (Fig. 3B). Such a difference between these two agonists has often been observed in many responses of activated neutrophils.

Activation of phosphatidylcholine synthesis by receptor-mediated agonists has been rarely considered. We show that fMLP triggers a typical cytidylyltransferase translocation (Fig. 4). In the case of PMA an additional activation of the membrane form of cytidylyltransferase occurs, which accounts for an increase of about 50% of total activity (Fig. 4C). The mechanisms leading to such activation remain to be established. Surprisingly, only a weak difference in membrane-bound cytidylyltransferase is observed after oleic acid treatment as compared to control, whereas oleic acid has been reported so far as the most potent inducer of cytidylyltransferase translocation (4). Since we could observe a net oleic acid-induced cytidylyltransferase translocation in undifferentiated HL-60 cells (37), but not in neutrophils, we can suggest that the fatty acid efficiency depends upon the stage of cell differentiation.

Among the regulators of cytidylyltransferase activity, diglyceride has been reported to control cytidylyltransferase translocation (5) . At variance with all the reports to date (4, 6) , the fatty acid has a weak effect on mass diglyceride formation (Fig. 5A). This is consistent with the slight cytidylyltransferase translocation to membranes. In contrast with growing cells (6) , oleic acid appears poorly metabolized into diglyceride in the human neutrophil. It should be noticed that the effect of oleic acid and PMA on the diglyceride generation in human neutrophil, i.e. in a differentiated cell, is exactly the opposite of what we observed previously in a tumor cell (43) .

We then questioned the origin of the diglyceride involved in cytidylyltransferase translocation. First, with fMLP as agonist, the absence of cytochalasin B blocks phosphatidylcholine breakdown at the phophatidic acid step (Fig. 6A), with no subsequent effect on phosphatidylcholine resynthesis and probably on cytidylyltransferase translocation. This result indicates that in vivo generation of an anionic phospholipid, such as phosphatidic acid, is not involved in cytidylyltransferase translocation. When phosphatidic acid is converted into diglyceride in the presence of cytochalasin B (Fig. 6B), then cytidylyltransferase translocation occurs. The alkaloid compound cytochalasin B, has been shown to release calcium from an intracellular pool (44) which might activate phosphatidate phosphatase (45) . The chemotactic peptide fMLP induces the formation both of phosphatidylinositol-derived and phosphatidylcholine-derived diglyceride, through phosphoinositide-phospholipase C (39) and phosphatidylcholine-phospholipase D/phosphatidate phosphatase pathways, respectively (9) . Another report (46) demonstrates that in the absence of cytochalasin B, fMLP induces a weak and transient formation of phosphoinositide-derived diglyceride, since no diglyceride can be recovered after 1-min of cell activation. Because no stimulation of [H]choline occurs in the absence of cytochalasin B (Fig. 3A), phosphoinositide-derived diglyceride could not be involved in cytidylyltransferase translocation in our cell model.

Second, an additional argument arises from the use of PMA, which has been shown to inhibit phosphoinositide phospholipase C (47) . Therefore diglyceride generated with PMA stimulation can only originate from phosphatidylcholine. We have confirmed that PMA inhibits phosphoinositide-derived diglyceride formation in neutrophils (Fig. 7A), and we demonstrate that the membrane-associated cytidylyltransferase parallels the kinetics of phosphatidylcholine-derived diglyceride. It is noteworthy that phosphatidylcholine-derived diglyceride is generated only after a 5-min lag time (Fig. 7B), the same lag time also being noticed in the time course of cytidylyltransferase association with membranes (Fig. 7C), in the mass increase of diglyceride (Fig. 5B), in the [H]choline incorporation (Fig. 3B) and in the phosphocholine decrease in the chase experiment (Fig. 2B, panel b). The results we obtained using a differential labeling of phosphoinositide and phosphatidylcholine strongly support the idea that PMA-induced cytidylyltransferase translocation is related to the generation of phosphatidylcholine-derived diglyceride only.

Third, the results obtained with the non-agonist compound oleic acid further strengthens a relationship between phospholipase D and cytidylyltransferase when cells are activated by agonists. The fatty acid could lead to diglyceride synthesis either by incorporation of oleic acid into monoglyceride (45) , or by activation of the phospholipase D/phosphatidate phosphatase pathway. Oleic acid has been shown to stimulate invitro phospholipase D from brain (48) , but only one report demonstrates invivo activation of the enzyme in hepatocytes (49) . In the neutrophil we demonstrate that oleic acid is unable to stimulate the phosphatidylcholine-phospholipase D, as assessed by the absence of phosphatidate or phosphatidylethanol formation (Fig. 6). We propose that only diglyceride formed through monoglyceride acylation plays a role in the slight cytidylyltransferase translocation induced by oleic acid.

To assess the point that phospholipase D activation is absolutely required for further phosphatidylcholine resynthesis, we have blocked fMLP-induced phospholipase D activation with the tyrosine kinase inhibitor tyrphostin (Fig. 8). Tyrphostin has no effect on the phosphoinositide-phospholipase C, which is activated with G proteins from the 7-helix fMLP receptor (8) , and therefore cannot prevent phosphoinositide-derived diglyceride. Also, tyrphostin does not affect cytidylyltransferase which is phosphorylated only on serine residues (5). Since cell treatment with this inhibitor effectively blocks both phosphatidylcholine degradation and resynthesis, this demonstrates that only phospholipase D-mediated phosphatidylcholine degradation is required for subsequent resynthesis of the phospholipid. As summarized in Fig. 9, diglyceride coming from phosphatidylcholine breakdown stimulates the reverse pathway by activating the rate-limiting enzyme. Activation of phosphatidylcholine turnover by the receptor-mediated agonist fMLP involves tyrosine kinases, whereas activation appears mediated by protein kinase C with the non-receptor agonist PMA, since the 4-O-methyl analog is ineffective. The effect of PMA triggering the phospholipase D/phosphatidate phosphatase pathway with subsequent enhancement on choline incorporation could explain the results of several reports showing a PMA-stimulated phosphatidylcholine synthesis, with a phorbol ester target being necessarily distinct of cytidylyltransferase (5, 6, 20) .


Figure 9: Relationship between cytidylyltransferase and phospholipase D activation in agonist-stimulated human neutrophils. Diglyceride coming from phosphatidylcholine breakdown activate the reverse pathway by promoting cytidylyltransferase translocation. The receptor-mediated agonist fMLP triggers a complete phosphatidylcholine cycle only when phosphatidic acid is converted into diglyceride, i.e. in the presence of cytochalasin B. In the absence of the alkaloid compound, phosphatidylcholine breakdown stops at the phosphatidic acid step, with no further effect on choline incorporation. Receptor-mediated activation of phosphatidylcholine cycle is mediated by tyrosine kinase-dependent phospholipase D stimulation, whereas the non-receptor agonist PMA appears to act through a protein kinase C-dependent phospholipase D activation. Oleic acid does not trigger a phosphatidylcholine cycle, but stimulates only the CDP-choline pathway.



Although the maximum activation of fMLP-induced phospholipase D is reached within 1 min (Fig. 6A and 8A), choline incorporation is maximum only after 15 min (Fig. 8B). This suggests many steps between the two processes of phosphatidylcholine breakdown and resynthesis. One of these could be that the choline pool used for phosphatidylcholine synthesis requires some time for labeling, before an effect of the agonist on choline incorporation could be noticed. This pool appears distinct from the one corresponding to the phospholipase D-mediated choline release. Effectively, about half of the choline released in Fig. 8A is recovered outside the cells (not shown). Such observations fit in with the scheme of George and co-workers (41) indicating that choline originating from phospholipid degradation is released out of the cell, whereas choline involved in synthesis is ``channeled'' up to the appropriate enzymes.

The human neutrophil appears, therefore, to be a good model for studying cytidylyltransferase activation in a differentiated cell, and for investigating the signaling events leading to the enzyme phosphorylation or dephosphorylation (50) . Potential sites for phosphorylation with mitogen-activated protein kinases have been recently identified in the enzyme sequence (51) . The human neutrophil which contains agonist-activable p40 and p42 isoforms (52) , should be a convenient model to investigate the relevance of cytidylyltransferase regulation with mitogen-activated protein kinases invivo.


FOOTNOTES

*
The part of work related to phospholipase D has been supported with a contract between INSERM and BAYER-PHARMA. The costs of publication of this article were defrayed in part by the payment of page charges. This article must therefore by hereby marked ``advertisement'' in accordance with 18 U.S.C. Section 1734 solely to indicate this fact.

§
Recipient of an allowance from BAYER-PHARMA.

To whom correspondence should be addressed. Tel.: 33-61-49-18-53; Fax: 33-61-49-67-49.

The abbreviations used are: PMA, phorbol 12-myristate 13-acetate; fMLP, formyl-methionyl-leucyl-phenylalanine; 4-O-methyl-PMA, phorbol 12-myristate 13-acetate 4-O-methyl ether; GPC, glycerophosphocholine.


REFERENCES
  1. Reinhold, S. L., Zimmerman, G. A., Prescott, S. M., and McIntyre, T. M.(1989) J. Biol. Chem. 264, 21652-21659 [Abstract/Free Full Text]
  2. Daniel, L., Waite, M., and Wykle, R. L.(1986) J. Biol. Chem. 261, 9128-9132 [Abstract/Free Full Text]
  3. Pelech, S. L., and Vance, D. E.(1989) Trends Biochem. Sci. 14, 28-30 [CrossRef]
  4. Vance, D. E.(1989) in Phosphatidylcholine Metabolism (Vance D. E., ed) pp. 225-239, CRC Press, Boca Raton, FL
  5. Tronchère, H., Record, M., Tercé, F., and Chap, H.(1994) Biochim. Biophys. Acta 1212, 137-151 [Medline] [Order article via Infotrieve]
  6. Utal, A. K., Jamil, M., and Vance, D. E.(1991) J. Biol. Chem. 266, 1405-1413
  7. Kiss, Z., Chattopadhyay, J., and Garanzregi, N.(1992) Arch. Biochem. Biophys. 296, 457-461 [Medline] [Order article via Infotrieve]
  8. Agwu, D. E., McPhail, L. C., Chabot, M. C., Daniel, L. W., Wykle, R. L., and McCall, C. E.(1989) J. Biol. Chem. 264, 1405-1413 [Abstract/Free Full Text]
  9. Gélas, P., Ribbes, G., Record, M., Tercé, F., and Chap, H. (1989) FEBS Lett. 251, 213-218 [CrossRef][Medline] [Order article via Infotrieve]
  10. Billah, M. M., Eckel, S., Mullmann, T. J., Egan, R. W., and Siegel, M. I.(1989) J. Biol. Chem. 264, 17069-17077 [Abstract/Free Full Text]
  11. Gélas, P., Von Tscharner, V., Record, M., Baggiolini, M., and Chap, H.(1992) Biochem. J. 287, 67-72 [Medline] [Order article via Infotrieve]
  12. Garcia Gil, M., Alonso, F., Sanchez-Crespo, M., and Mato, J. M.(1981) Biochem. Biophys. Res. Commun. 101, 740-748 [Medline] [Order article via Infotrieve]
  13. Bomalaski, J. S., Dundee, D., Brophy, L., and Clark, M. A.(1990) J. Leukocyte Biol. 47, 1-12 [Abstract]
  14. Pike, M. C., Kredich, N. M., and Snyderman, M.(1980) Cell 20, 373-379 [Medline] [Order article via Infotrieve]
  15. Mato, J. M., Pencev, D., Vasanthakumar, G., Schiffmann, E., and Pastan, I.(1983) Proc. Natl. Acad. Sci. U. S. A. 80, 1929-1932 [Abstract]
  16. Garcia Gil, M., Alonso, F., Alvarez Chiva, V., Sanchez Crespo, M., and Mato, M.(1982) Biochem. J. 206, 67-72 [Medline] [Order article via Infotrieve]
  17. Pelech, S. L., Pritchard, P. H., Brindley, D. N., and Vance E. D. (1983) J. Biol. Chem. 258, 6782-6787 [Abstract/Free Full Text]
  18. Cornell, R., and Vance, D. E.(1987) Biochim. Biophys. Acta 919, 37-48 [Medline] [Order article via Infotrieve]
  19. Guy, G. R., and Murray, A. W.(1982) Cancer Res. 42, 1980-1985 [Abstract]
  20. Cook, H. W., Byers, D. M., Palmer, F. B. St. C., and Spence, M. W. (1989) J. Biol. Chem. 264, 2746-2752 [Abstract/Free Full Text]
  21. Kolesnick, R. N.(1987) J. Biol. Chem. 262, 14525-14530 [Abstract/Free Full Text]
  22. Watkins, J. D., and Kent, C.(1991) J. Biol. Chem. 266, 21113-21117 [Abstract/Free Full Text]
  23. Billah, M. M., and Anthes, J. C.(1990) Biochem. J. 269, 281-291 [Medline] [Order article via Infotrieve]
  24. Strum, J. C., Nixon, A. B., Daniel, L. W., and Wykle, R. L.(1993) Biochim. Biophys. Acta 1169, 25-29 [Medline] [Order article via Infotrieve]
  25. Thompson, N. T., Bonser, R. W., and Garland, L.(1991) Trends Pharmacol. Sci. 12, 404-409 [CrossRef][Medline] [Order article via Infotrieve]
  26. Record, M., Laharrague, P., Fillola, G., Thomas, J., Ribbes, G., Fontan, P., Chap, H., Corberand, J., and Douste-Blazy, L.(1985) Biochim. Biophys. Acta 819, 1-9 [Medline] [Order article via Infotrieve]
  27. Bligh, E. G., and Dyer, W. J.(1959) Can. J. Biochem. Physiol. 37, 911-917
  28. Vance, D. E., Pelech, S. L., and Choy, P. C.(1981) Methods Enzymol. 71, 576-581 [Medline] [Order article via Infotrieve]
  29. Pelech, S. L., Paddan, M. B., and Vance, D. E.(1984) Biochim. Biophys. Acta 795, 447-451 [Medline] [Order article via Infotrieve]
  30. Tercé, F., Record, M., Ribbes, G., Chap, H., and Douste-Blazy, L. (1988) J. Biol. Chem. 263, 3142-3149 [Abstract/Free Full Text]
  31. Wright, T. M., Rangan, L. A., Shin, H. S., and Raben, D. M.(1988) J. Biol. Chem. 263, 9374-9380 [Abstract/Free Full Text]
  32. Preiss, J., Loomis, C. R., Bishop, W. R., Stein, R., Niedel, J. E., and Bell, R. M.(1986) J. Biol. Chem. 261, 8597-8600 [Abstract/Free Full Text]
  33. Olson, S., Bowman, E. P., and Lambeth, D. J.(1991) J. Biol. Chem. 266, 17236-17242 [Abstract/Free Full Text]
  34. Lowry, O. H., Rosebrough, N. J., Farr, A. L., and Randall, R. J.(1951) J. Biol. Chem. 193, 265-275 [Free Full Text]
  35. Wroblewski, F., and La Due, J. S.(1955) Proc. Soc. Exp. Biol. Med. 90, 210-215
  36. Record, M., Bes, J. C., Chap, H., and Douste-Blazy, L.(1982) Biochim. Biophys. Acta 688, 57-65 [Medline] [Order article via Infotrieve]
  37. Tronchère, H., Tercé, F., Record, M., Ribbes, G., and Chap, H.(1991) Biochem. Biophys. Res. Commun. 176, 157-165 [Medline] [Order article via Infotrieve]
  38. Cabot, M. C., Welsh, C. J., Zhang, Z. C., Cao, H. T., Chabott, H., and Lebowitz, M.(1988) Biochim. Biophys. Acta 959, 46-57 [Medline] [Order article via Infotrieve]
  39. Ohta, H., Okajima, F., and Ui, M.(1985) J. Biol. Chem. 260, 15771-15780 [Abstract/Free Full Text]
  40. Tercé, F., Record, M., Tronchère, H., Ribbes, G., and Chap, H.(1991) Biochim. Biophys. Acta 1084, 69-77 [Medline] [Order article via Infotrieve]
  41. George, T. P., Cook, H. W., Byers, D., Palmer, F. B. St. C., and Spence, M. W.(1991) J. Biol. Chem. 266, 12419-12423 [Abstract/Free Full Text]
  42. Bentley, J. K., and Reed, P. W.(1981) Biochim. Biophys. Acta 678, 238-244 [Medline] [Order article via Infotrieve]
  43. Tronchère, H., Tercé, F., Record, M., and Chap, H.(1993) Biochem. J. 293, 739-744 [Medline] [Order article via Infotrieve]
  44. Treves, S., Di Virgilio, F., Vaselli, G. M., and Pozzan, T.(1987) Exp. Cell. Res. 168, 285-298 [Medline] [Order article via Infotrieve]
  45. Jamil, H., Utal, A. K., and Vance, D. E.(1992) J. Biol. Chem. 267, 1752-1760 [Abstract/Free Full Text]
  46. Honeycutt, P. J., and Niedel, J. E.(1986) J. Biol. Chem. 261, 15900-15905 [Abstract/Free Full Text]
  47. Tyagi, S. R., Tamura, M., Burnham, D. N., and Lambeth, J. D.(1988) J. Biol. Chem. 263, 13191-13198 [Abstract/Free Full Text]
  48. Chalifour, R., and Kanfer, J. N.(1982) J. Neurochem. 39, 299-305 [Medline] [Order article via Infotrieve]
  49. Siddiqui, R. A., and Exton, J. H.(1992) Eur. J. Biochem. 210, 601-607 [Abstract]
  50. Houweling, M., Jamil, H., Hatch, G. M., and Vance, D. E.(1994) J. Biol. Chem. 269, 7544-7551 [Abstract/Free Full Text]
  51. MacDonald, J. I. S., and Kent, C.(1994) J. Biol. Chem. 269, 10529-10537 [Abstract/Free Full Text]
  52. Torres, M., Hall, F. L., and O'Neill, K.(1993) J. Immunol. 150, 1563-1578 [Abstract/Free Full Text]

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