platelets (71 % of total activity) and was released into the extracellular ... The secretion of sPLA, correlated with the release of ATP. sPLA,- ..... (Tokyct) IOS, 903 -.
Eur. J. Biochem. 216, 169-175 (1993) 0 FEBS 1993
Secretory phospholipase A, is not required for arachidonic acid liberation during platelet activation Carine MOUNIER', Ahmad FAILI', B. Boris VARGAFTIG', Cassian BON' and Mohamed HATMI,
'
Unit6 des Venins, Institut Pasteur, Paris, France * Unit6 de Pharmacologie Cellulaire, Unite AssociCe Institut Pasteur-INSERM 285, Institut Pasteur, Paris, France (Received April 6May 21, 1993) - EJB 93 0502/6
The subcellular localization of secretory phospholipase A, (sPLA,) and cytosolic phospholipase A, (cPLA,) in resting and activated platelets, and their involvement in arachidonic acid liberation during platelet activation, were studied. The amounts of sPLA, and cPLA, recovered were not modified during platelet activation. sPLA, was mainly associated with the organelles of resting platelets (71 % of total activity) and was released into the extracellular medium during cell activation (60% of total activity), whereas the majority of cPLA, was localized in the cytosol of resting and activated platelets. The secretion of sPLA, correlated with the release of ATP. sPLA,-depleted platelets aggregated as much as control platelets and produced similar amounts of thromboxane B, upon thrombin activation. These results indicate that sPLA, is not involved in the liberation of arachidonic acid during platelet activation.
Phospholipases A, (PLA,) hydrolyse the sn-2-acyl bond of sn-3-phosphoglycerides and may therefore release arachidonic acid and lyso-platelet-activating factor (lyso-PAF), which are the precursors of the inflammatory lipid mediators, eicosanoids and PAF, respectively [l -51. Two mammalian PLA2, which may be involved in arachidonic acid liberation, have been described. Secretory PLA, (sPLA,), is released upon in-vitro or in-vivo activation of platelets from different species and has been purified [6-91 as a 14-kDa protein containing 124 amino acids, from which the gene has been cloned [lo, 111. sPLA, requires millimolar levels of Ca2+ and a high pH for activity and thus shares similarities with extracellular mammalian and venom PLA,. More recently, cytosolic PLA, (cPLA,) has been described in different mammalian species and various cells, including platelets [12171. It is a protein of 749 amino acids with a theorical molecular mass of 85.2 kDa [18], with a marked specificity towards phospholipids, such as phosphatidylcholine, phosphatidylserine, phosphatidylinositol and phosphatidylethanolamine, which bear arachidonic acid in the sn-2 position [4], and is activated at physiological concentrations of intracellular Ca2+(0.3 - 1 yM). In resting platelets, sPLA, is localized in Q granules, in agreement with its secretory nature [7, 19, 201, whereas cPLA, is cytosolic. After activation, cPLA, is partially translocated to the membrane in the presence of Ca'' [13, 21Correspondence to C. Bon, Unit6 des Venins, Institut Pasteur, 25 rue du Dr Roux, F-75724 Paris Cedex 15, France Abbreviations. Pam(PyrDec)GroPMe, 1-palmitoyl-2-(10-pyrenyldecanoyl)-sn-glycero-3-phosphomethane;Pam(PyrDec)GroPCho, 1 -palmitoyl-2-(1O-pyrenyldecanoyl)-sn-glycero-3-phosphocholine ; Pam(PyrDec)GroPGro, 1-palmitoyl-2-(1 O-pyrenyldecan0yl)-~sn-glycero-3-phosphoglycerol ; cPLA,, cytosolic phospholipase A,; PAF, platelet-activating factor; sPLA,, secretory phospholipase A,; TxB,, thromboxane B,. Enzyme. Phospholipase A, (EC 3.1.1.4).
24). The specific functions of both enzymes remain unclear, and in particular, their involvement in the release of arachidonic acid has not been determined. This is of a particular interest, because the low intracellular levels of free arachidonic acid suggest that PLA, activation in platelets represents a rate-limiting step in the process of lipid-mediator synthesis [1, 31. Due to its arachidonic acid specificity, its low Ca2+requirement and its subcellular localization, it is likely that cPLA, is involved in arachidonic acid release. However, recent studies indicated that the sPLA, may participate in eicosanoid formation following hydrolysis of the cellular phospholipids of activated inflammatory cells, i.e., vascular smooth-muscle cells, granulocytes and mast cells [25-271. To determine the respective role of these enzymes, we investigated whether sPLA, is involved in the production of eicosanoids during platelet activation. We first established the subcellular localization of both types of PLA, in resting and activated platelets, and studied the release of sPLA,. In addition, we compared the thromboxane B, (TxB,) formation in sPLA,-depleted and control platelets upon stimulation by thrombin. Our results show that sPLA, is not involved in TxB, formation during platelet activation and support the hypothesis that cPLA, is responsible for eicosanoid formation.
MATERIALS AND METHODS Materials Platelet-activating factor (PAF) was obtained from Bachem and bovine thrombin from Hoffmann-La-Roche. Fibrinogen (grade L) from Kabi was treated with diisopropyl fluorophosphate to inactivate pro-coagulant contaminants. Prostacyclin, EGTA, EDTA, bovine serum albumin (fraction V), and fatty-acid-free bovine serum albumin, were from Sigma. Antibodies, thromboxane B, (TxB,) used as standard, and radiolabelled ligands for TxB, determinations were ob-
170 tained from the URIA (Institut Pasteur). ATP was from glycero-3-phosphocholineand 4 nmol diacylglycerol as subSigma, luciferine and luciferase from Lumac, and the Brad- strate vesicles [34]. After introducing the sample into the ford reagent from Bio-Rad. /?-nicotinamide adenine dinucleo- incubation medium (200 p1 final volume), the reaction protide was from Sigma and 4-nitrophenyl-P-D-glucopyrano- ceeded at 37 "C for 30 min and was terminated with the addisiduronic acid from Merck. Radiolabelled substrate, l-pal- tion of chlorofordmethanol (1:2, by vol.). Lipid extraction mitoyl- 2 - ['4C]arachidonoyl- sn - glycero - 3 - phosphocholine was performed according to Bligh and Dyer [35], the organic (52 Ci/mol) was from NEN Research Products. Thin-layer- phases were dried and redissolved in 50 pl chlorofordmethchromatography plates (0.2 mm) and standard phospholipids anol (1:1, by vol.). Samples were spotted on silica plates and were from Merck. Fluorescent substrates, 1-palmitoyl-2-(10- chromatographed in chloroform/methanol/water/acetic acid pyrenyldecanoyl)-sn-glycero-3-phosphomethane [Pam(Pyr- (65 :43 :3 :1, by vol.). Authentic non-labelled phospholipids Dec)GroPMe] and 1-palmitoyl-2-(10-pyrenyldecanoy1)-sn- were used as standards and visualized by iodine vapor. The glycero-3-phosphocholine [Pam(PyrDec)-GroPCho] were areas containing phosphatidyl and free arachidonic acid were from Interchim, and 1-palmitoyl-2-(l0-pyrenyldecanoyl)- cut out, transferred into vials, and analysed for radioactivity sn-glycero-3-phosphoglycerol [Pam(PyrDec)GroPGro) was by liquid scintillation. sPLA, should not have any activity on this substrate since we established that this enzyme does not from KSV. hydrolyse PC.
Preparation of rabbit platelets Male rabbits (HY/CR strain Charles River) of 3-4 kg were bled from the central artery of the ear in 5 mM EDTA as an anti-coagulant. Platelet-rich plasma was collected after blood centrifugation at 375 g for 20 min at 20"C, and centrifuged for 15 min at 1800 g and 20°C. The pellet was washed twice with Tyrode's buffer (137 mM NaC1, 2.7 mM KCl, 11.9mM NaHCO,, 0.42 mM NaH,PO,, 1 mM MgCl,, 5.6 mM glucose, 0.35% bovine serum albumin) adjusted to pH 6.5 and completed with 0.2 mM EGTA and 1 mM MgC1,. Platelets were then resuspended in Tyrode's buffer, pH 7.4, and counted automatically. Platelet lysates were obtained with a 10-min treatment of the platelet suspension by ultrasound at 40 W (type 20 Sonimasse sonicator).
Subcellular fractionation Platelet fractionation was performed as described previously [28], with minor modifications. Briefly, platelets (2 ml fraction, 2 X 10' cells/pl) were incubated for 5 min at 37°C with thrombin 0.25 U/ml (activated cells), or with its solvent (resting cells), then centrifuged for 5 min at 7800 g. The supernatant fraction (S) was collected and the pellet was resuspended in 2 ml fresh Tyrode's buffer, pH 7.4. Resuspended platelets were lyzed by ultrasonication at 40 W for 4 X 5 s, then centrifuged at 19000 g, for 25 rnin at 4°C. The pellet (organelle fraction, 0) was resuspended in Tyrode's buffer and stored, and the supernatant was centrifuged at 100000 g for 60 min. The supernatant (cytosolic fraction, C) was removed and the pellet (membrane fraction, M) was gently dispersed in Tyrode's buffer. Each fraction of resting or stimulated platelets was assayed for the following markers : lactate dehydrogenase, /?-glucuronidase and proteins as described ([29-311, respectively), and for sPLA, and cPLA, activities.
Phospholipase A, assays
Platelet aggregation Aggregation was monitored by light transmission with an Icare aggregometer with continuous stirring at 1100 rpm. Platelets at 5 X lo5 cells/pl were incubated for 2 rnin with 2 mM CaCI,, and aggregation was followed for 3 min after the addition of the agonist, except in the kinetic experiments. Appropriate siliconized glass tubes were used for aggregation. Agonists were PAF (10 nM, plus fibrinogen 0.28 mg/ ml, final concentration) and thrombin at the indicated concentrations. Each sample was decanted in eppendorf tubes with a glass pipette then immediately centrifuged for 2 min at 7800 g and 4°C in order to stop the release reaction. The supernatant was then used for the assays of TxB,, lactate dehydrogenase, ATP, and sPLA, activity. The presence of the sPLA, in glass tubes as well as in the glass pipette may provoke a loss of sPLA, activity due to non specific absorption to the walls, which may explain why in such experimental conditions only 33% of the enzyme was released.
TxB, radioimmunoassay TxB, was determined with a slightly modified [33] radioimmunoassay [36]. ATP release The ATP content was determined 3 rnin after stimulation by the bioluminescence method (luciferine-luciferase) [37] using a Pico-ATP device from Jobin et Yvon.
Preparation of sPLA,-depleted platelets Platelet suspension (3 ml) containing 5 X lo5 cells/pl were stimulated at room temperature with 50 nM PAF under stirring with a Teflon magnetic stirrer. After 3 min, 10 pM prostacyclin and 0.4 mM EDTA were added, and the platelets were gently stirred for 30 rnin to allow their disaggregation. The platelet suspension was then centrifuged for 10 rnin at 20°C and 1800 g, and the pellet was resuspended in fresh Tyrode's buffer, pH 7.4. Immediately before adding prostacyclin and EDTA, an aliquot was collected, centrifuged at 7800 g, and the supernatant was used to assay lactate dehydrogenase, ATP and sPLA,.
sPLA, activity was assayed with Pam(PyrDec)GroPMe as a substrate, according to the method of Radvanyi et al. [32] with minor modifications [33]. Two other substrates, Pam(PyrDec)GroPGro and Pam(PyrDec)GroPCho, were used to examine the substrate specificity of the sPLA,. It is expected that cPLA, should not cleave any of these substrates, because of its specificity for phospholipids with Electron microscopy arachidonic acid at the sn-2 position. The cPLA, activity was measured with a radioassay using Control and sPLA,-depleted platelets at 5 X lo5 cells/pl a mixture of 6 nmol l-palmit0y1-2-[~~C]arachidonoyl-sn-were prepared for electronmicrographs, as already described
171 6
A
S
O
M
C
T
S
O
M
C
T
Fig. 1. Subcellular localization of sPLA, and cPLA,. Platelets were fractionated as described in Materials and Methods and subcellular fractions were tested for P-glucuronidase (A), lactate dehydrogenase (B), sPLA, (C) and cPLA, (D) activities. Fraction S corresponds to extracellular medium, C to cytosol, 0 to organelles, M to membrane, and T to platelet lysate. Closed columns correspond to resting platelets and open columns to platelets activated with thrombin (0.25 U/ml). Values are the means rt SEM of 3-7 separate experiments.
[38]. Briefly, platelets were fixed with 3% glutaraldehyde in 0.1 M sodium phosphate, pH 7.3 for 1 h at 4°C. The pellet was then post-fixed with 1% OsO, in the same buffer, rinsed and dehydrated with acetone before embedding in Epon. Thin sections were obtained on diamond knife, contrasted with uranyl acetate and lead nitrate, and examined with a Philips TEMISTEM microscope operating at 80 kV.
Statistical analysis Statistical significance between treated and control samples was performed using a Student's t-test for unpaired data.
RESULTS AND DISCUSSION Subcellular localization of sPLA, and cPLA, in resting and thrombin-stimulated platelets Fig. 1A and B show that the organelle fraction contained the p-glucuronidase activity, whereas the cytosolic fraction contained most of the lactate-dehydrogenase activity. The recovery of the enzymes after fractionation was in the range 94-110% (data not shown). As described by Kim et al. [39], sPLA, and cPLA, were present in platelets. The recovery of sPLA,, evaluated with the fluorescence assay, was 95% in resting and 100% in activated cells. sPLA, was predominantly associated with the organelle fraction in resting cells and was released into the extracellular medium during platelet activation by thrombin (Fig. 1C). Thus, only 6% of the total activity was found in the supernatant of resting cells, compared to 60% for activated cells. The increase of sPLA, activity observed in the supernatant during stimulation correlated with its decrease in the organelle fraction (71% in resting cells versus 20% in activated cells). No increase in the total activity of sPLA, between resting and activated cells was noted, indicating that the activity of sPLA, was not modified during platelet activation. Fig. 1D shows that 67% of the cPLA, activity was localized in the cytosolic fractions of resting and activated cells. It has been shown that the cPLA, of macrophages is associ-
ated in part with the membranes in the presence of 0.11 pM CaZ+[21], and translocated to the membranes of stimulated macrophages in the presence of 235-450 nM Caz+ [24]. Translocation of cPLA, has been described in rat brain synaptosomes but only at high Ca2+concentrations [22], and also correlated with the differentiation of human monocytic tumour-cell lines [23]. In contrast, we failed to observe translocation of cPLA, after platelet activation. It is nevertheless likely that such translocation has occured and that failure to note it results from the dissociation of the enzyme from the membrane, particularly because of the low Ca2+ concentrations used during cell fractionation. An increase in the total activity of cPLA, after fractionation, with recovery of 146% in resting cells and 136% in stimulated cells, was observed. This probably results from a fractionation artifact, such as an alteration of the enzyme environment, due to the separation of the membrane phospholipids from the cytosolic fraction [40]. However, we failed to note an increase in cPLA, activity upon platelet activation, whereas such increase was described during activation of mesangial cells [41], macrophages [42], and monocytes [43] after incubations of 10-30 min, compared to less than 5 min in our conditions. This may be due to different mechanisms in the different cell types. A transient increase in cPLA,-mRNA levels has been detected in stimulated monocytes 15 min after stimulation [43]. Such a mechanism cannot be involved in platelets since they are anucleated and rapidly activated. However, platelets might lack a specific kinase or kinase-regulatory components, which are responsible for the activation of cPLA, by phosphorylation in macrophages 1421 and ovary cells [44].
Characteristics of the sPLA, To examine the substrate specificity of sPLA, we used three fluorescent substrates containing the same fatty acid at the sn-1 position and a 10-pyrene group at the sn-2 position, but different polar heads, glycerophosphomethane and glycerophosphoglycerol, which are negatively charged, or glycerophosphocholine, which is neutral at physiological pH. Table 1 shows that sPLA, displayed a marked specificity for
172 Table 1. Substrate specificity and pH dependence of sPLA, activity. sPLA, activity was measured in subcellular fractions from resting and thrombin-activated platelets (as given in legend of Fig. 1) with the fluorometric assay using Pam(PyrDec)GroPMe (PA), Pam(PyrDec)GroPGro (PG) and Pam(PyrDec)GroPCho (PC) as substrates. Values are the means ? SEM of three separate experiments. ~~
Platelet type
Subcellular fraction
sPLA, activity with PG at
PA at
PC at
pH 7.4
pH 6.5
pH 7.4
PH 8
132 3 170 + 35 lo+ 1 462 5 230 + 49 592 7 282 3 87 -C 15
142 1 190 + 15 12+ 1 53t 4 260 i 20 69? 5 33+ 3 1002 8
pH 7.4
nmol . min-’ . lo9 cells -’ Resting platelets
S 0 M C
64 ? 10 850 ? 130 525 8 230 2 35
Activated platelets
S 0 M C
1150 + 170 300? 45 1 4 0 5 20 440 + 70
8 +- 0.4 100 t 5 6 t 1.5 28 +- 1.5
140 t 7 36 5 2 17 ? 1 52 ? 3
Ca2+ concentration (mM)
Fig. 2. Caz+-dependenceof sPLA, activity. The sPLA, activity contained in the organelle fraction of resting platelets (0),or in the supernatant of activated platelets (O),was determined with a fluorescence assay in the presence of 1 mM EGTA and 0.5-10 mM CaCI,. Free Ca2+ was calculated as described by Durham [48]. Values are the means 2 SEM of three separate experiments.
a
t-
a
negatively charged substrates, and was inactive towards Pam(PyrDec)GroPCho. In addition, Pam(PyrDec)GroPGro was a better substrate than Pam(PyrDec)GroPMe. The same substrate specificity was found for all fractions, before and after platelet activation, suggesting the presence of a single enzyme which was not modified by activation. This conclusion was confirmed by analysing the sPLA, activity of each fraction at different pH (Table 1). Finally, sPLA, was only active in the presence of millimolar concentrations of Ca” (Fig. 2). Clearly, platelets contain a single sPLA, which is not affected by cell activation.
Dose/response and kinetic curves of sPLA, release during platelet activation by thrombin The concentrations of thrombin required to induce the secretion of sPLA,, aggregation and ATP release were similar (Fig. 3). Fig. 4 further shows that ATP and sPLA, release were almost concomitant, 50% release being reached after 27 s and 32 s, respectively. Aggregation was delayed compared to ATP and sPLA, release, since 50% aggregation was recorded only after 53 s (Fig. 4). In a previous study [ 3 3 ] , we have shown that PAF, collagen and the stable endoperoxide analog compound U46619, like thrombin, also induced sPLA, release, indicating that the release of sPLA, is an indicator of platelet activation.
4
c
Thrombin ( U / m l )
Fig. 3. Dose/response curve for platelet activation by thrombin. Platelets were incubated for 2 min with CaC1, (2 mM), then stimulated with different concentrations of thrombin. Aggregation (A), ATP (B) and sPLAz activity (C) were determined after 3 min. Values are the means ? SEM of five separate experiments.
TxB, formation by sPLA,-depleted platelets Platelets were stimulated with 50 nM PAF during 3 min then disaggregated with the addition of prostacyclin and EDTA. After centrifugation, the supernatant containing the
173
60 40
-
20
-
Offdl 0
-
r
0 100
&
50
100
150
200
5 0
100
150
200
150
200
lc
0
50
100
Time
(5)
Fig.4. Kinetics of aggregation, and of ATP and sPLA, release during platelet activation. Platelets were incubated for 2 min with CaC1, (2 mM) then activated with 0.25 U/mlthrombin. Aggregation (A), ATP (B) and sPLA, (C) were determined after various time intervals. Values are the means -t SEM of five separate experiments.
sPLA, was removed, and the sPLA,-depleted platelets were resuspended in Tyrode's buffer. Depleted platelets were not damaged, since the leakage of lactate dehydrogenase activity was below lo%, and platelet electronmicrograph examination showed undamaged cells and a marked degranulation (Fig. 5). As mentioned for thrombin (Fig. l), platelet activation induced by PAF did not increase the total sPLA, activity (331 5 76nmol . min-' . 1Oj cells-' in activated platelets versus 292 f 75 nmol . min-' . lo9 cells-' in control platelets). PAF-activated platelets had released 90% of their ATP content, and 70% of their sPLA, content (234 t 58 nmol . min-' . lo9 cells-' in the supernatant compared to 331 f 76 nmol . min. ' . lo" cells-' in the lysate of activated platelets). The sPLA, activity which remained associated with platelets after their activation (less than 30%) may result either from an incomplete degranulation, or more probably from the absorption of the released enzyme onto the plasma membrane, as suggested by Kurihara et al. [25]. Interestingly, sPLA,-depleted platelets could be further activated. This differs from the results of Horigome et al. [7], who reported that the release of a significant fraction of sPLA, was accompagnied by irreversible aggregation. We obtained similar aggregation with sPLA,-depleted and control platelets, in response to thrombin (Fig. 6A). As expected, the sPLA,-depleted platelets did not release ATP when stimulated (Fig. 6C), and only a small amount of sPLA,
Fig. 5. Transmission electron microscopy views. (A) control platelets; (B) sPLA,-depleted platelets. Horizontal bars represent 1 pm.
(Fig. 6D). In the absence of activation, the sPLA,-depleted platelets showed higher sPLA, release than untreated control platelets (Fig. 6D). The presence of sPLA, in the supernatant may therefore reflect the resorption of sPLA, adsorbed to the membrane during the depletion step. More importantly, the sPLA,-depleted platelets produced similar amounts of TxB, as control platelets (Fig. 6B), indicating that the liberation of arachidonic acid was not affected, and thus that sPLA, is not involved in the production of eicosanoids. In conclusion, we failed to observe an increase in the total activity of sPLA, and cPLA, during platelet activation, which is compatible with the very short delays of platelet activation. Release of sPLA, is as much an index of platelet activation as aggregation and ATP release. More interestingly, our observations show that sPLA, is not implicated in the formation of arachidonic acid during platelet activation, indicating the exclusive involvement of cPLA,, as suggested for other cells [12-14, 16, 24). Indeed, the participation of cPLA, in arachidonic acid liberation during activation of Chinese hamster ovary cells was recently shown [44]. The exact function of platelet sPLA, remains unclear, and a role in the amplification of the inflammatory process, in analogy with other released platelet products, may be suggested, since
174 8
A
C -T
._ .E
-.8
.-m
20
_I
0
control
lrealed
control
'I
treated
140 120
40
20
control
trealed
control
**
A lrealed
Fig. 6. Comparison between the activation of control and sPLA,-depleted platelets. Control (untreated) and sPLA,-depleted (treated) platelets were incubated for 2 min with CaC1, (2 mM), then activated by thrombin (0.25 U/ml). Aggregation (A), TxB, formation (B), ATP (C) and sPLA, activity (D) were determined 3 rnin after saline (a)or thrombin (B) addition. Values are the means t SEM of fair separate experiments. Statistical significance (**, P < 0.01 and ***, P < 0.001) compared to control platelets.
its presence in body fluids correlates with the degree of inflammation [45-471. We are greatly indebted to Dr Pierre Gounon and Ms MarieChristine PrCvost (Station Centrale de Microscopie Electronique, Institut Pasteur) for electronmicrographs. We thank Prof. Claude Jacquemin for his helpful comments during writing of the manuscript. This research was supported in part by funds from the Direction des Recherches et Etudes Techniques.
REFERENCES 1. Chang, J., Musser, J. H. & McGregor, H. (1987) Biochem. Pharmacol.36, 2429-2436. 2. Kroll, M. H. & Schafer, A. I. (1989) Blood 74, 1181-1195. 3. Lapetina, E. G. (1982) Trends Pharmacol. Sci. 3, 115-118. 4. Axelrod, J. (1990) Biochem. Soc. Trans. 18, 503-507. 5. Yamada, K., Okano, Y., Miura, K. & Nozawa, Y. (1987) Biochem. J. 247, 995-999. 6. Mizushima, H., Kudo, I., Horigome, K., Murakami, M., Hayakawa, M., Kim, D. K., Kondo, E., Tomita, M. & Inoue, K. (1989) J. Biochem. (Tokyo) 105, 520-525. 7. Horigome, K., Hayakawa, M., Inoue, K. & Nojima, S. (1987) J. Biochem. (Tokyo) 101, 53-61. 8. Kramer, R. M., Hession, C., Johansen, B., Hayes, G., McGray, P., Chow, E. P., Tizard, R. & Pepinsky, R. B. (1989) J. Biol. Chem. 264, 5768-5775. 9. Murakami, M., Kudo, I. & Inoue K. (1989) Biochim. Biophys. Acta 1005, 270-276. 10. Kusunoki, C., Satoh, S., Kobayashi, M. & Niwa, M. (1990) Biochim. Biophys. Acta 1087, 95-97. 11. Komada, M., Kudo, I. & Inoue K. (1990) Biochem. Biophys. Res. Commun. 168, 1059-1065. 12. Clark, J. D., Milona, N. & Knopf, J. L. (1990) Proc. Nut1 Acad. Sci. USA 87, 7708-7712. 13. Clark, J. D., Lin, L. L., Kriz, R. W., Ramesha, C. S., Sultzman, L. A., Lin, A. Y., Milona, N. & Knopf, J. L. (1991) Cell 65, 1043-1051. 14. Sharp, J. D., White, D. L., Chiou, X. G., Goodson, T., Gamboa, G. C., McClure, D., Burgett, S., Hoskins, J., Skatrud, P. L., Sportsman, J. R., Becker, G. W., Kang, L. H., Roberts, E. F. & Kramer, R. M. (1991) J. B i d . Chem. 266, 14850-14853.
15. Kim, D. K., Kudo, I. & Inoue, K. (1991) Biochim. Biophys. Acta 1083, 80-88. 16. Takayama, K., Kudo, I., Kim, D. K., Nagata, K., Nozawa, Y. & Inoue, K. (1991) FEBS Lett. 282, 326-330. 17. Kim, D. K., Suh, P. G. & Ryu, S. H. (1991) Biochem. Biophys. Res. Commun. 174, 189-196. 18. Tremblay, N. M., Nicholson, D., Potier, M. & Weech, P. K. (1992) Biochem. Biophys. Res. Commun. 183, 121-127. 19. Horigome, K., Hayakawa, M., Inoue, K. & Nojima, S. (1987) J. Biochem. (Tokyo) 101, 625-631. 20. Van den Bosch, H., Aarsman, A. J. & Verkleij, A. J. (1989) in Leukotrienes and prostanoids in health and disease, (Zor, U., Naor, Z. & Danon, A,, eds) vol3, pp. 257-261, Kargel, Basel. 21. Krause, H., Dieter, P., Schulze-Specking, A., Ballhorn, A. & Decker, K. (1991) Eul: J. Biochem. 199, 355-359. 22. Yoshihara, Y. & Watanabe, Y. (1990) Biochem. Biophys. Res. Commun. 170, 484-490. 23. Rehfeldt, W., Hass, R. & Goppelt-Stmebe, M. (1991) Biochem. J . 276, 631-636. 24. Channon, J. Y. & Leslie, C. C. (1990) J. Biol. Chem. 265, 5409-5413. 25. Kurihara, H., Nakano, T., Takasu, N. & Arita, H. (1991) Biochim. Biophys. Acta 1082, 285-292. 26. Hara, S., Kudo, I. & Inoue, K. (1991) J. Biochem. (Tokyo) 110, 163-165. 27. Murakami, M., Kudo, I. & Inoue, K. (1991) FEBS Lett. 294, 247- 251. 28. Le Peuch, C. J., Le Peuch, D. A. M., Katz, S., Demaille, J. G., Hincke, M. T., Bredoux, R., Enouf, J., Levy-Toledano, S. & Caen, J. (1983) Biochim. Biophys. Acta 731, 456-464. 29. Johnson, M. K. (1960) Biochem. J. 77, 610-618. 30. Talalay, P., Fishman, W. H. & Huggins, C. (1946) J. Biol. Chem. 166, 757-762. 31. Bradford, M. M. (1976) Anal. Biochem. 72, 248-254. 32. Radvanyi, F., Jordan, L., Russo-Marie, F. & Bon, C. (1989) Anal. Biochem. 177, 103- 109. 33. Mounier, C., Hatmi, M., Faili, A., Bon, C. & Vargaftig, B. B. (1993) J. Pharmacol. Exp. Ther: 264, 1460-1467. 34. Kramer, R. M., Ghecani, G. C. & Deykin, D. (1987) Biochem. J. 248, 779-783. 35. Bligh, E. G. & Dyer, W. J. (1959) J. Biochem. Physiol. 37, 911-915.
175 36. Sors, H., Pradelles, P., Dray, F., Rigaud, M., Maclouf, J. & Bernard, P. (1978) Prostaglandins 16, 277-290. 37. Holmsen, H., Storm, E. & Day, H. J. (1972) Anal. Biochem. 46, 489-501. 38. Pottu-Boumendil, J. (1989) Microscopic Electronique, yrincipes et mCthodes de pripamtion, INSERM ed., 222. 39. Kim, D. K., Kudo, I., Fujimori, Y.,Mizushima, H., Masuda, M., Kikuchi, R., Ikizawa, K. & Inoue, K. (1990) J. Biochem. (Tokyct) IOS, 903 -906. 40. Kramer, R. M., Checani, G. C., Deykin, A,, Pritzker, C. R. & Deykin, D. (1986) Biochim. Biophys. Acta 878, 394-403. 41. Gronich, J. H., Bonventre, J. V. & Nemenoff. R. A. (1988) J. Biol. Chem. 263. 16645-16651
42. Wijkander, J. & Sundler, R. (1992) FEBS Lett. 311, 299-301. 43. Nakamura, T., Lin, L.-L., Kharbanda, S., Knopf, J. & Kufe, D. (1992) EMBO J. 11, 4917-4922. 44. Lin, L-L., Lin, A. Y. & Knopf, J. L. (1992) Proc. Nut1 Acc~d. Sci. USA 89, 6147-6151. 45. Tesson, F., RuffiC, C., Hidi, R., Silva, P., Vazeux, G., Vargaftig, B. B. & Bon, C. (1993) Eul: J. Pharm, in the press. 46. Vadas, P. & Pruzanski, W. (1986) Lab. Invest. 55, 391 -404. 41. Pruzanski, W. & Vadas, P. (1991) Immunol. Today 12j 143146. 48. Durham, A. C. H. (1983) Cell Calcium 4, 33-46.
