Eur. J. Immunol. 1979. 9: 409-415
Jean Michel Mencia-Huerta and Jacques Benveniste INSERM U 131, Clamart
Release of platelet-activating factor from macrophages
409
Platelet-activating factor and macrophages I. Evidence for the release from rat and mouse peritoneal macrophages and not from mastocytes* Platelet-activating factor (PAF) is a phospholipid mediator of anaphylaxis, released from basophils of several mammalian species, that aggregates platelets and releases their vasoactive amines. The ionophore A 2 3 187 induced the release of PAF from rat and mouse peritoneal cells, a mixed cell population that was fractionated using 5-15 % Ficoll gradients and adherence to plastic petri dishes. PAF was associated with large, acid phosphatase-containing, adherent mononuclear cells. Mastocytes did not release PAF but released histamine by the action of ionophore or 48/80; they could not be held responsible for inactivation of PAF or inhibition of the PAF-induced platelet aggregation. These data indicate that, besides blood basophils, peritoneal macrophages are a likely source for PAF, a result that adds a new important function to the macrophage: aggregation of platelets and liberation of their inflammatory and vasoactive substances.
1 Introduction
2 Materials and methods
Platelet-activating factor (PAF) is a phospholipid mediator that causes aggregation of platelets and triggers the release reaction [l-31. Liberation by PAF of potent phlogistic and vasoactive substances from platelets constitutes an amplifying mechanism that is instrumental in immune complex deposition in acute serum sickness in rabbits [4, 51. Numerous lines of evidence indicate that PAF is released from circulating basophils. In immunized rabbits, PAF has been obtained from leukocytes in the presence of the specific antigen, o r anti-IgE antiserum [2]. The presence of histamine-containing cells has been well correlated with the release of a soluble factor [6], now known to be identical with PAF. We have presented ultrastructural evidence of rabbit platelets aggregating around degranulating basophils [2]. In addition, a factor bearing some resemblance to PAF has been obtained from a pure basophil preparation from human leukemic blood [7].
2.1 Buffers
Activators known to interact with mastocyte membrane receptors [8, 91, such as 48/80, C 5 a and neutrophil cationic proteins, were capable of releasing PAF from human leukocytes and human and rabbit tissues [lo]. Accordingly, with others [11], we suspected mastocytes as a likely source for PAF. In order to examine this hypothesis, we have studied peritoneal cells (PC) from rats and mice using various cell separation procedures. In this study, we will show that, contrary to our expectations, we have recovered PAF from macrophage-like cells and not from purified mastocytes. [I 22431
* This work was supported by INSERM: AT No. 77-79, and DGRST: ACNO.77-7-1409. Correspondence: Jacques Benveniste, INSERM U 131, 32 rue des Carnets, F-92140 Clamart, France Abbreviations: P A F Platelet-activating factor PC: Peritoneal cells HEPES: N-2-Hydroxyethylpiperazine-N-2-ethanesulfonic acid BSA. Bovine serum albumin HBCM: HEPES buffer containing calcium and magnesium ions HB: HEPES buffer SRS-A: Slow-reacting substance of anaphylaxis ECF-A: Eosinophil chemotactic factor of anaphylaxis
0Verlag Chemie, GmbH, D-6940 Weinheim, 1979
RPMI 1640 (Flow Labs., Irvine, Scotland) was prepared by dissolving 10.46 g of powder in 1 liter of distilled water and M (final) N-2-hydroxythen buffered to pH 6.8 with 5 X ethylpiperazine-N’-2-ethanesulfonicacid (HEPES) (Calbiochem, Los Angeles, CA). In some experiments, 2.5 g of bovine serum albumin (BSA) were added to RPMI (RPMI-BSA) and the p H adjusted to 7.4 with 0.1 N NaOH. A HEPES-buffered medium containing calcium and magnesium ions (HBCM) was prepared as described in [12]:for 1 liter, we used 4.77 g HEPES, 0.37 g KCl, 8.18 g NaC1, 1.00 g glucose, 0.20g CaCl, X 6H,O and 0.10 g MgCl, X 6 H z 0 . The same buffer was prepared without calcium and magnesium (HB). When 2.5 gAiter BSA were added to HBCM and HB, they were designated as HBCM-BSA and HB-BSA.
2.2 Preparation of PC One to 3-month-old Wistar rats and C57BL/6 mice of both sexes were used. PC were collected according to Bloom et al. [13]. Briefly, RPMI (10 ml for rats or 3 ml for mice) was injected into the peritoneal cavity of exsanguinated and etherkilled animals. After abdominal massage, the peritoneum was opened and the injected fluid collected with siliconized Pasteur pipettes. The cells were washed twice by centrifugation (200 X g, 5 min) and resuspended in suitable media, generally HBCM. The cells were stained by mixing 10 pl of the cell suspension with 90 p1 of 0.1% toluidine blue solution in 10% formaldehyde, SO% ethanol and 1 % acetic acid. This solution selectively stains the mastocyte granules [ 141. Differential cell counts were performed on smears stained with May-Griinwald-Giemsa solution. Histochemical staining of peroxidase-positive cells was performed as described by Preud’homme et al. [15]. In some experiments, PC were collected 48 h to 1 week after an intraperitoneal injection of distilled water [16]. Such cell preparations were devoid of mastocytes. 0014-2980/79/0S05-0409$02.S0/0
410
Eur. J . Immunol. 1979. 9: 409-415
J. M. Mencia-Huerta and J. Benveniste
2.3 Fractionation of PC 2.3.1 Ficoll gradients A stock solution of 15% (w/v) of Ficoll (Pharmacia, Uppsala, Sweden) in distilled water was buffered with 20mM HEPES and the pH adusted to 6.8 with 1N NaOH. A solution of 5% (w/v) Ficoll was obtained by addition of 2 volumes of HB to 1 volume of 15 % Ficoll solution. Gradients were prepared in 14-ml cellulose nitrate tubes (14 mm X 94 mm) with a gradient former (Isco, Lincoln, NE) using 6.5 ml of 5% and 6.5 ml of 15% Ficoll solutions. One X lo7 PC in 500 pl of RPMI were layered on the top of the gradient and centrifuged at 200 X g for 3 rnin at room temperature. Fractions of approximatively 850 pl each were collected with a fraction collector (Auto densi-flow, Buchler, Fort Lee, NJ). The fractions were then diluted 2-fold with HB-BSA, and the cells were pelleted by centrifugation at 200 X g for 15 min, washed twice with HB-BSA, sedimented by centrifugation (200 X g, 5 min) and finally resuspended in 0.4 ml HBCM-BSA or RPMI-BSA. This method was preferred to that of Sullivan et al. [17] or Orange et al. [12] which in our experiments did not yield mononuclear cell-rich fractions totally free of mastocytes.
5 min, the cells were resuspended in 200 p1 of RPMI, preheated for 10 rnin at 3 7 ° C and 1 pg/ml of ovalbumin then added. The reaction was stopped 5 min later by addition of 50 pl of the formaldehyde-containing staining solution. Cells were examined microscopically 30 min after staining and the number of degranulated vs. intact mastocytes assessed. This technique was performed either on whole PC or on Ficoll gradient-purified mastocytes.
2.6 Assay procedures PAF activity was determined in an aggregometer apparatus (Icare, Marseille, France) by aggregation of 5 X lo6 to 7 X lo6 washed rabbit platelets in 300 pl Tyrode's buffer in the presence of 5 p~ indomethacin (Sigma). It was expressed in arbitrary units as measured by the relative chart distance for the aggregation produced by 10 plof PAF-containingsolution. The preparation of platelets and the composition of buffers have been described in detail elsewhere [2]. Histamine was measured by an automatic spectrofluorometric method [ 191, and acid phosphatase with p-nitrophenyl phosphate as substrate [20] after lysis of cells with Triton X-100. Cell viability was assessed before and after each experiment by the trypan blue exclusion method.
2.3.2 Cell adherence One X lo7 cells in RPMI-BSA (2 ml) were plated in 3.5 cm diameter treated plastic petri dishes (3001 Flow) and incubated at 37 " C for 6 h. The dishes were thoroughly washed with RPMI-BSA after 2,4 and 6 h. Nonadherent cells were pooled, pelletted by centrifugation (200 X g, 5 min) and resuspended in 1 ml of RPMI-BSA.
2.4 Procedure for cell incubation The following substances were used on whole or purified PC: ovalbumin (Sigma Chemical Co., St. Louis, MO); compound 48/80 (Sigma) was dissolved in suitable media immediately before use; a stock solution of 5 mg/ml ionophore A 23187 (Elli Lilly Labs., Indianapolis, IN) was prepared in dimethyl sulfoxide and stored at - 20°C. It was diluted with saline buffer to suitable concentrations immediately before use. Equivalent dilutions of dimethyl sulfoxide were added as control. Reactions were carried out at 37 'C for 60 min, unless otherwise indicated, and were stopped by centrifugation (200 X g, 5 min) or by addition of 2.5 pl/ml of 0.2 M EDTA. In the case of plastic-adherent cells, release experiments were performed directly in petri dishes, by addition of activating substances; the incubation media were collected, centrifuged and stored. In some cases, maximal releasable PAF was obtained by overnight incubation of the cells at room temperature in HBCM-BSA at p H 10.6 [3]. All supernatants were stored at - 20 " C until assay of the released substances.
2.5 Passive sensitizationof rat PC Passive sensitization was performed according to A. ProuvostDanon et al. [18]. Briefly, 2 X l o 6 PC in 200 pl RPMI were incubated for 30 rnin at 3 7 ° C under agitation with various dilutions of IgE-rich mouse anti-ovalbumin antiserum with a titer > 1: 10000 (a gift of A. Prouvost-Danon). After 3 washings with RPMI alternated with centrifugations at 200 X g for
2.7 Characterization of PAF Aggregation of platelets by PAF was not influenced by indomethacin. In addition, two criteria to distinguish PAF from other possible aggregating agents were applied. PAF-containing solutions were incubated with phospholipase A, (Boehringer, Mannheim, FRG), as described [21], or chromatographed on silica-gel plates. To perform the thin-layer chromatography, 3 volumes of absolute ethanol wereaddedto 1volume of the PAF solution. After 25 rnin under agitation, the tubes were centrifuged at 1500 X g for 30 min. The supematants were evaporated to dryness, and PAF was recovered by addition of 200 pl of ethanol, 50 to 100 pl of which were layered on silica-gel plates 60F254 (Merck Darmstadt, FRG). The plates were developed in chloroform: methanol: water (70 :35 :7) and either stained by exposition to iodine vapor or divided into 10 X 10 mm squares. The gel of each square was recovered by scraping, and PAF activity was eluted with 100 pl of HBCM-BSA.
3 Results 3.1 Effect of the ionophore and 48/80 on unfractionated rat PC Upon addition of the ionophore A23187 or 48/80 to 5 X lo6 PC in 1 ml of HBCM-BSA, release of PAF was time-dependent and maximal after 40 min for the ionophore, 60 rnin for 48/80. The 60-rnin interval was chosen for all following exM periments. The release of PAF was abolished by 5 X EDTA or by incubation at 4 "C. By contrast with 48/80 which yielded low and variable amounts of PAF, the ionophore induced high releases from all PC populations. Cell viability was not affected, as less than 2 % of cells lost their ability to exclude trypan blue after 1 h incubation with or without ionophore or 48/80. By contrast, with PAF, histamine release was
Eur. J. Immunol. 1979. 9: 409-415
Release of platelet-activating factor from macrophages
already maximal at 5 min and at the same level with either the ionophore or 48/80 (Fig. 1).
A
We have studied the effect of increasing concentrations of ionophore on the release of PAF and histamine and mastocyte degranulation. The release of PAF was nearly maximal with 0.5 pg/ml ionophore when mastocyte degranulation was less than 14%, in contrast to histamine which closely followed degranulation (Fig. 2). Also, comparable amounts of PAF were obtained from a variety of PC showing large differences in their mastocyte numbers. Finally, PC from rats depleted of mastocytes by a previous injection of distilled water released PAF at the same level as PC from uninjected animals (Fig. 3).
%
41 1
A
A
h
2 A-
mean * s d
A A A
42.4 t 4 . 2 A
A A A
A
Units
1001
r50
Figure 3. Comparison of the release of PAF with the number of mastocytes in rat PC from different animals. (r = 0.134; not significant). PAF was obtained from 5 X 106 PC in 1 ml HBCM-BSA by incubation with 1 p g h l ionophore for 60 min and expressed as shown in Fig. 1. Mastocytes were counted microscopically afterstainingby toluidine blue. (A) PC from normal rats. ( A t )PC from rats injected with distilled water 3-7 days before experiments.
-
n
100%
al
10
20
30
40
50
60
90
T i m e (min)
Figure I . Release of histamine (---) and PAF (-) from 5 X 106rat PC in 1 ml HBCM-BSA by 1 pg/ml ionophore (A) or 1 pg/ml48/80 ( 0 ) .(0) incubation with HBCM-BSA alone, or at 4"C, or with 5x M EDTA. Histamine is expressed as the percentage of the amount found in supernatants over the total cell content (8.0 -t 3.0pg). PAF is expressed in arbitrary units, as described in Sect. 2.6. The 40-min incubation of PC with the ionophore yielded as much releasable PAF as the exposure of cells to alkaline pH. Data represent the mean f 1 standard deviation of 5 experiments.
a
r
1jBiTT-j
T u b e number
Figure 4 . Cumulative percentage of different cell types of rat PC after fractionation on 5-15 % Ficoll gradient. (A) Small lymphocytes; (B) macrophages, large lymphocytes and monocytes; (C) mastocytes; (---) concentrations of Ficoll. This figure is representative of 20 cell separation experiments. In this and all following figures, gradient fractions are numbered from top (left, abscissa) to bottom (right, abscissa).
3.2 Distribution of rat PC after Ficoll gradient separation J
I
0.05
jmmj
0.1
0.5
1
5 pglml
lonophore
Figure 2. Degranulation of mastocytes (0--0),release of PAF (A-A) and histamine (&--A) after incubation of 5 X 106 rat PC with increasing amounts of ionophore for 60 min. PAF and histamine are expressed as shown in Fig. 1. Data represent the mean k 1 standard deviation of 3 experiments (degranulation and histamine release) and of 5 experiments for release of PAF.
The distribution of PC, shown in Fig. 4, is expressed in cumulative percentages for each cell type. This method produced a good separation of mastocytes which sedimented at the bottom of the gradient, from macrophages and lymphocytes, which were more buoyant. To some extent, macrophages and large lymphocytes could be separated from small lymphocytes. Cell recovery was 70 to 80%, and the purity of mastocyte populations was between 95 and 99%. Fractions containing lymphocytes and macrophages were totally devoid of mastocytes.
412
J. M. Mencia-Huerta and J. Benveniste
Monocytes, identified as peroxidase-positive cells, accounted for 5 % of the total populations and were recovered in fractions 4 and 5 . In contrast to the results of others [22], neutrophils were very rare in our preparations (0.96 k 0.63% in 5 experiments).
3.3 Release of PAF from fractionated rat PC Exposure to alkaline p H of Ficoll fractions yielded PAF from the upper region of the gradient, corresponding to mononuclear cells, i.e. large lymphocytes, monocytes andmacrophages. PAF was associated with acid phosphatase-containing cells and was neither obtained from the top fractions corresponding to small lymphocytes nor from the lower region corresponding to mastocytes. Only the latter fractions yielded histamine (Fig. 5 ) . When fractions were incubated with 1 y g h l ionophore, PAF was again obtained only from the upper region of the gradient (Fig. 6). Cell fractions were also incubated with 1 pg/ml48/80. Neither PAF nor any other aggregating activity was detected in any region of the gradient (10 experiments), whereas release of histamine (and degranulation of masto-
Eur. J. ImmunoI. 1979. 9: 409-415
cytes) occurred as expected in the mastocyte-containing tubes (3 experiments) (Fig. 6).
3.4 Separation of mouse PC Separation of mouse PC was rather poor as compared to rats. Nevertheless, PAF coincided with acid phosphatase activity in a region where few mastocytes were present (Fig. 7, bottom). The amount of PAF and acid phosphatase activity obtained from mastocyte-depleted mice was comparable to that of normal mice (Fig. 7, top).
3.5 Effect of Ficoll fractionation on rat cell populations The following experiments were performed to check whether centrifugation in Ficoll was capable of causing cell damage. (a) The loss in cell viability, as assessed by trypan blue exclusion, was less than 2 % after the separation and washing procedures; (b) the amount of PAF released by the ionophore from unfractionated PC was identical to that of the same population, either separated and pooled back to the original amount of cells or incubated 15 min in 15% Ficoll solution and then washed; (c) mastocytes which sedimented in the 12 to 15% Ficoll fractions did not degranulate spontaneously but degranulated and released their histamine upon addition of
Figure 5. Presence of PAF (-0) (mean 4 1 standard deviation of 8 experiments), acid phosphatase (0-0) and histamine (A-A) in Ficoll gradient-fractionated rat PC. PAF was released at alkaline pH, acid phosphatase was measured after lysis of cells by Triton X-100 and histamine after boiling the cell preparations for 5 min.
Units
10-6
501
1 . 12
p%q Figure 6. Release of PAF from Ficoll gradient-fractionated rat PC or 48/80 (0); release of histamine by by 1 kg/ml ionophore (A-A) 1 pg/ml 48/80 ( C - - 0 ) . Each point represents the mean f 1 standard deviation of 6 experiments for the ionophore, 10 experiments (4WXO-PAF) and 3 experiments (48/80-histamine).
Ix
14
Tube n u m b t i
Figure 7. PAF (0-0) and acid phosphatase (0-0) from Ficoll gradient-fractionated mouse PC. Top: PC harvested 3 days after intraperitoneal injection of distilled water. Bottom: untreated mice. (W-W) total cell counts. Vertical bars represent mastocyte number. These data are representative of 5 experiments.
Eur. J . Immunol. 1979. 9: 409-415
Release of platelet-activating factor from macrophages
1 pg/ml of 48/80 (Table 1 and Fig. 6); (d) PC passively sensitized with mouse anti-ovalbumin antiserum were fractionated and incubated with 1 pg of ovalbumin. Mastocytes degranulated to the same degree, before and after passage through the gradient (Table 1). In some experiments, passive sensitizations with the anti-ovalbumin antiserum were performed after passage of mastocytes through the Ficoll gradient. Upon addition of the antigen, they degranulated as well as mastocytes from unfractionated PC (results not shown). Table 1. Effect of the separation procedure on rat peritoneal mastocytes
Hi\tamine release by 4x 80;'' Ikgrsnulation of nia\ttryte\"' Spontaneous 13) 4X. XI) O\alhuinind'
Before gradient separation
After gradient separation
63.5 t 4 9 '
66.3 t 6.0
7.7 t 3.0 70.3 t 5.0 74.0 3.3
7.1 3.9 78.9 t -1.0 73.0 f 0.8
*
*
a) Mastocytes in 200 p.l of RPMI were incubated 5 min at 37 "C with 1 pg/ml 48/80. Histamine release was assessed as described in Sect. 2.6 and expressed as percentage of total content. b) Data represent the mean t 1 standard deviation of 5 experiments. c) Counts were performed 30 min after addition of 50 p1 of the stainig solution to 200 p.l of the mastocyte suspension and degranulation expressed as percentage of degranulated vs. intact mastocytes. d) PC were sensitized by incubation for 30 min at 3 7 ° C with IgE-rich mouse anti-ovalbumin antiserum (final dilution 1 : 500). They were then exposed to 1 p g h l ovalbumin for 5 min and mastocyte degradation assessed microscopically.
3.6 Role of mastocytesin the release or inhibition of PAF Purified rat mastocytes from Ficoll gradients were pooled and incubated with 1 pg/ml of 48/80 or ionophore, or after passive sensitization with IgE-rich anti-ovalbumin antiserum, with 1 pg/ml ovalbumin. No release of PAF was obtained. To determine whether the lack of PAF release could in fact be due to the degradation of PAF or to inhibition of PAF-induced aggregation of platelets, purified mastocytes were sonicated or lysed with distilled water. The cell lysates were then incubated
4 13
with PAF (approximatively 30 units/l0 PI). No loss in PAF activity was found after 1 h at 37 "C. Also, we could not detect any inhibitory effect of the cell lysates or of supernatants from 48/80-degranulated purified mastocytes on the PAF-induced aggregation of platelets, therefore the presence of an inhibitor of platelet aggregation can be ruled out. To determine whether spontaneous release of PAF from mastocytes or its inactivation occurred during the passage of the cells through the Ficoll solution, we prepared gradients containing either 0.25% BSA as a carrier of PAF [2] or PAF (30 units/l0 pl). After deposition of PC, centrifugation and recovery of fractions, no spontaneous release of PAF or loss of activity could be demonstrated (3 experiments). 3.7 Fractionation of PC by adherence PAF was obtained from adherent rat cells by action of the ionophore or alkaline pH (Table 2); 48/80 failed to induce the release of PAF from adherent or nonadherent cells. A small amount of PAF was obtained from nonadherent cells by action of the ionophore. In an attempt to show whether PAF release occurred during the adherence of macrophage's, media from petri dishes were extracted with 3 volumes of ethanol, centrifuged, evaporated to dryness and then resuspended in 100 ~1 of HBCM-BSA. No PAF activity was recovered.
3.8 Characterization of PAF PAF obtained from macrophages before and after Ficoll gradient, migrated as a lyso-phosphatidylcholine on silica-gel thinlayer chromatography (Rf : 0.35), between sphingomyelin and phosphatidylcholine. This chromatographic behavior was identical to that of PAF from blood leukocytes (Fig. 8). Arachidonic acid, another platelet-aggregating agent, migrated with the solvent front. Three thousand units/ml of PAF from macrophages were completely inactivated, as well as PAF from leukocytes, by phospholipase A, from Naja-naja or Crotalus. PAF from macrophages or leukocytes aggregated platelets in the presence of 5 pM indomethacin, a concentration that totally suppressed an equivalent aggregation by 10 p~ arachidonic acid. +-solvent front ---------------------or
Table 2. Release of PAF from adherent and nonadherent cellsa)
.. .. H
Treatment
Adherent cells
Nonadherent cells w
Nil 3X/XO
lonophorc pH 10.0
0 Traces 35 t 9' 39 7
+
0 Traces 12 5-1 7 t6
.o
4-
Rf 0.35
. . . . . . 1
a) Rat PC were allowed to adhere on plastic petri dishes for 6 h. Nonadherent cells were recovered by washing at 2 , 4 and 6 h. 1 pg/ml 48/80 or ionophore was then added. Supernatants (1 ml) were collected after 1 h incubation at 37 "C. In some experiments, release of PAF was obtained by exposure to pH 10.6, as described [3]. b) Results are expressed in arbitrary units as the mean ? 1 standard deviation of 6 experiments.
H
2
3
4
5
6
;
m Figure 8. Diagrammatic representation of PAF on silica-gel thinlayer chromatography. (1) PAF from pig blood leukocytes; (2) PAF from rat PC before and (3) after Ficoll gradients; (4) sphingomyelin; (5) phosphatidylcholine; (6) lyso-phosphatidylcholine; (7) arachidonic acid. Solvent = chloroform :methanol : water (70 :35 : 7).
414
Eur. J. Immunol. 1979. 9: 409-415
J. M. Mencia-Huerta and J. Benveniste
4 Discussion The release of PAF was obtained by the action of the ionophore A 23187 or compound 48/80 on rat and mouse PC, a mixed population of lymphocytes, macrophages and mastocytes. The release of PAF by the ionophore was not correlated with the number of mastocytes. It was identical in mastocytedepleted PC and in whole PC population. PC were centrifuged on Ficoll gradients, and fractions containing lymphocytes and mononuclear cells (including macrophages) released PAF, but 95 % pure mastocyte preparations were inactive. PAF was present in fractions containing acid phosphatase, a marker for macrophages [23], and in adherent cells, but not in fractions containing histamine (only present in mastocytes) and in small amounts in nonadherent cell populations. The liberation of PAF by the ionophore A 23 187 was an active process which was inhibited by EDTA or low temperatures. Besides the ionophore, 48/80 or antigen also induced degranulation of purified mastocytes and the release of histamine, but failed to liberate PAF. Negative results were also observed by incubation of mastocytes at pH 10.6, a procedure which is very efficient in releasing PAF from blood leukocytes [3] or lymphomononuclear-rich PC fractions. The lack of PAF release from mastocytes could not be explained by its spontaneous release or degradation during the separation procedure. All these results exclude mastocytes and indicate that macrophages are a likely source for PAF. Besides histamine, 48/80 released PAF from whole PC - although much less than ionophore or alkaline pH - but not from fractionated PC containing either lymphocytes and macrophages, or mastocytes. This indicates that mastocytes could intervene in the release of PAF, possibly through an interaction with macrophages :degranulation of mastocytes and release of histamine (and other mediators) in a first step, then, in a second step, phagocytosis of the mastocyte granules by macrophages. This hypothesis is supported by the fact that we have obtained the release of PAF along with lysosomal enzymes, during phagocytonis of zymosan particles - another argument in favor of the macrophage origin of PAF - and observed the presence of mastocyte granules in macrophage phagosomes after incubation of whole PC with 48/80 ([24], and manuscript in preparation). A two-step process analogous to our observations could explain the well-documented IgEdependent release of slow-reacting substance of anaphylaxis (SRS-A) 1251 and eosinophil-chemotactic factor of anaphylaxis (EFC-A) [26]. Our results do not allow us to determine whether all adherent cells were involved in the release of PAF. Adherent cells are not only macrophages [27], and we cannot exclude the possibility that another cell than the macrophage could also generate PAF. These restrictions are common in all situations where cell purification could not be performed to a sufficient degree. Our macrophage-derived PAF appeared to be physicochemically identical with PAF from human, rabbit, hog and horse leukocytes, as assessed by the currently available methods of identification. The finding of a platelet-activating agent raises the question of its biochemical nature. A substance with platelet-activating activity, not further identified, was obtained from a 100% pure human basophil preparation [7]. PAF was released from human buffy coat cells with anti-IgE antiserum, C5 a or neutrophil cationic proteins, well-known activators of cells of the mastocyte-basophil series [lo], and we have confirmed (Benveniste and Camussi, unpublished results) its iden-
tity with PAF from rabbit leukocytes. Besides in blood, PAF has been described as originating from various tissues [ 10, 11, 281. PAF was not precisely identified [ l l , 281, and we could not ascertain the nature of the factor obtained from lung and kidney in the experiments reported [lo]. We know now that the finding in media from immunologically stimulated cells or organs of a platelet-activating and/or releasing substance is not a valid criterion to affirm the presence of PAF, as previously defined [21]. Other aggregating agents can be found in bathing media from lungs and kidneys (Benveniste et al., manuscript in preparation); this is the case with arachidonic acid which mimics the effect of PAF on platelets but is inhibited by indomethacin. It remains, that to date no experimental data are available to confirm the existence of different molecular species of the 1-lyso-phosphatidylcholine with platelet-aggregating activity [21]. However, until the complete molecular structure of PAF is elucidated, the identity of these PAF from leukocytes or macrophages of various origins cannot be strictly defined. Nevertheless, given the similitude of their characteristics, any existing differences should probably be minor.
Our present data are not contradictory to the well-accepted basophil origin of PAF. However, they necessitate a reconsideration of the pathophysiological role of PAF (and other mediators): far from being specific of the cells involved in the mechanisms of immediate hypersensitivity, it may be one of the numerous mediators belonging to the cells of the phagocytic group. PAF was originally described in basophils from immunized rabbits after challenge with specific antigen or anti-IgE [2]. In contrast to our present results, this release was very rapid (less than 5 min), and a factor probably identical to PAF was well correlated with histamine release 161. More evidence was provided by electron microscopical studies where platelets were shown to aggregate around degranulated basophils [2]. We have discussed the facts in favor of the release of PAF from human basophils. Our findings of the macrophage origin of PAF is reminiscent of the problem of the various origins attributed to SRS-A and ECF-A, other mediators of immediate hypersensitivity [25, 261. SRS-A was first associated with the release of histamine [29, 301 and with the IgE-mastocyte system [25]. It was later obtained from polymorphonuclear leukocytes [311 and from rat peritoneal cells, regardless of the presence of mastocytes [32]. The mononuclear cell origin of SRS-A, upon stimulation by the ionophore, has recently been ascertained [22]. In this case also, mastocytes failed to release SRS-A. By contrast, SRS-A was released from purified mastocytes upon stimulation by the ionophore A23187 [33]. ECF-A has been found in polymorphonuclear leukocytes [34]. It is therefore likely that the mediators of immediate hypersensitivity, SRS-A, ECF-A and PAF, can be released by a variety of cell types under various stimuli. Our present results add a new function to the macrophage, aggregation of platelets and release of their vasoactive amines. A cooperation between macrophages and platelets has recently been described in tumoral cell growth inhibition [35]. Yet, the different mechanisms of release and the potential pathophysiological implication of the generation of PAF by macrophages remain to be elucidated.
Received August 31,1978.
IgG-binding lymphocytes control the expression of receptors for IgM
Eur. J. Immunol. 1979. 9: 415-420
5 References 1 Siraganian, R. P. and Osler, A. G., J. Immunol. 1971. 106: 1244. 2 Benveniste, J., Henson, P. M. and Cochrane, C. G., J. Exp. Med. 1972.136: 1356. 3 Benveniste, J., Nature 1974. 249: 581. 4 Henson, P. M. and Cochrane, C. G., J. Exp. Med. 1971.133: 554. 5 Benveniste, J., Egido, J. and Gutierrez-Millet, V., Clin. Exp. Immunol. 1976. 26: 449. 6 Siraganian, R. P. andOsler, A. G., J. Immunol. 1971. 106: 1252. 7 Lewis, R. A., Goetzl, E. J., Wasserman, S. I., Valone, F. H., Rubin, R. H. and Austen, K. F., J. Immunol. 1975. 114: 87. 8 Morrison, D. C., Roser, J. F. and Cochrane, C. G., Int. Arch. Allergy Appl. Immunol. 1975. 49: 172. 9 Ranadive, N. S. and Cochrane, C. G., J. Exp. Med. 1968. 128: 605. 10 Camussi, G., Mencia-Huerta, J. M. and Benveniste, J., Immunology 1977.33: 523. 11 Kravis, T. C. and Henson, P. M., J. Immunol. 1975.115: 1677. 12 Orange,R. P.andMoore, E. G., J.lmmunol.1976.117:2191. 13 Bloom, G. D., Fredholm, B. and Maegermark, O., Acta Physiol. Scand. 1967. 71: 270. 14 Mota, I . and Dias da Silva, W., Nature 1950.186: 245. 15 Preud'homme, J. L. and Flandrin, G., J. Immunol. 1974. 113: 1650. 16 Facette, W., J. Exp. Med. 1954. I00t217. 17 Sullivan, T., Parker, K. L., Eisen, S. A. and Parker, C. W., 1. immunol. 1975.114: 1473. 18 Prouvost-Danon, A,, Wyczolkowska, J., Binaghi, R. and Abadie, A., Immunology 1975.29: 151.
Christiane SamarutO and Jean-Pierre Revillard Laboratoire d'Immunologie, INSERM U-80 and C.N.R.S. ERA 782, Lyon
415
19 Ruff, F., Saindelle, A., Dutripon, E. and Parrot, J. L., Nature 1967.214: 279. 20 Linhardt, K. and Walter, K., in Bergmeyer, H.-U. (Eds.), Methods of Enzymatic Analysis, Academic Press, New York 1965, p. 776. 21 Benveniste, J., Le Couedic, J. P., Polonsky, J. and Tence, M., Nature 1977.269: 170. 22 Bach, M. K. and Brashler, J. R., J. Immunol. 1978.120: 998. 23 Cohn, Z. A. and Benson, R., J. Exp. Med. 1965.121: 153. 24 Benveniste, J. and Mencia-Huerta, J. M., Fed. Proc. 1978. 37: 1554. 25 Orange, R. P., Stechschulte, P. J. and Austen, K. F., J. Immunol. 1970.105: 1087. 26 Kay, A. B. and Austen, K. F., J. Immunol. 1971.107: 899. 27 Nathan, C. F., Asofsky, R. and Terry, W. D., J. Immunol. 1977. 118: 1612. 28 Kravis,T. C. andHenson, P. M., J. Immunol. 1977. 118: 1569. 29 Kellaway, C. H. and Trethewie, E. R., Quart. J. Exp. Physiol. 1940. 30: 121. 30 Schild, J . O., Hawkins, D. F., Mongar, J. L. and Herxheimer, J., Lancet 1951. i: 376. 31 Conroy, M. C., Orange, R. P. and Lichtenstein, L. M., J. Immunol. 1976.116: 1677. 32 Bach, M. K. and Brashler, J. R., J. Immunol. 1974.113: 2040. 33 Yecies, L. D., Wedner, H. J., Johnson, S. M. and Parker, C. W., Fed. Proc. 1978. 37: 1667. 34 Czametzki, B., Konig, W. and Lichtenstein, M. L., Nature 1975. 258: 725. 35 Johnson, R. J., Pastemack, G. R. and Shin, M. S., J. Immunol. 1977.118:494.
Human T lymphocyte receptors for IgM: control by IgG-binding lymphocytes* The capacity of human peripheral blood lymphocytes to bind antigen-IgG or antigenIgM-antibody complexes was investigated using a rosette technique with ox erythrocytes (E) coated with rabbit IgG (AG)or IgM (AM)antibodies. EAMrosette formation was achieved only in suspensions pre-incubated for 24 h at 37 "C. Addition of either EAMor EAG complexes to the culture medium was shown to prevent the formation of EAM rosettes. The inhibition was reversible, it was not due to trace IgM contaminants in the IgG antibody fraction. It was not observed when lymphocytes depleted of EAG-rosetting cells were incubated with EAG complexes. Inhibition of the expression of lymphocyte receptors for IgM can be regarded as a consequence of the modulation of surface receptors for IgG and involves an interaction between the two lymphocyte subsets bearing surface receptors for IgG and IgM, respectively. However, these experiments do not exclude the possibility that a few cells which bind EAG may lose their receptors by modulation and then express a receptor for IgM.
[I 22341
* C0ntractsD.G.R.S.T. 77.7.1381 andA.T.P. INSERM78-151. AttachCe de recherche (INSERM)
Correspondence: Jean-Pierre Revillard, HBpital E. Herriot, Pavillon P, F-69374 Lyon Cedex 2, France Abbreviations: E Ox erythrocytes EAG: Ox erythrocytes coated with rabbit IgG antibodies EA,: Ox erythrocytes coated with rabbit IgM antibodies FcyR. Lymphocyte receptor for IgG FcpR Lymphocyte receptor for IgM RFC: Rosette-forming cells TG: T lymphocytes bearing FcyR TM: T lymphocytes bearing FcpR 0 Verlag Chemie, GmbH, D-6940 Weinheim, 1979
1 Introduction Numerous cell types, including B, T and nulllymphocytes, bear readily detectable surface receptors (FcyR) for the Fc portion of aggregated o r antigen-bound IgG (reviewed in [l]).Receptors for the Fc part of native or antigen-bound IgM (FcpR) have also been detected on subsets of T [2, 31 and B cells [4, 51. FcyR and FcpR are most often expressed on different cells. Hence, two distinct subpopulations of human T cells have been defined by the presence of FcyR or FcpR and designated as TG and TM,respectively [6].These two subsets were reported to differ by several morphological [7] and 0014-2980/79/0505-0415$02.50/0
Jean Michel Mencia-Huerta and Jacques Benveniste INSERM U 131, Clamart
Release of platelet-activating factor from macrophages
409
Platelet-activating factor and macrophages I. Evidence for the release from rat and mouse peritoneal macrophages and not from mastocytes* Platelet-activating factor (PAF) is a phospholipid mediator of anaphylaxis, released from basophils of several mammalian species, that aggregates platelets and releases their vasoactive amines. The ionophore A 2 3 187 induced the release of PAF from rat and mouse peritoneal cells, a mixed cell population that was fractionated using 5-15 % Ficoll gradients and adherence to plastic petri dishes. PAF was associated with large, acid phosphatase-containing, adherent mononuclear cells. Mastocytes did not release PAF but released histamine by the action of ionophore or 48/80; they could not be held responsible for inactivation of PAF or inhibition of the PAF-induced platelet aggregation. These data indicate that, besides blood basophils, peritoneal macrophages are a likely source for PAF, a result that adds a new important function to the macrophage: aggregation of platelets and liberation of their inflammatory and vasoactive substances.
1 Introduction
2 Materials and methods
Platelet-activating factor (PAF) is a phospholipid mediator that causes aggregation of platelets and triggers the release reaction [l-31. Liberation by PAF of potent phlogistic and vasoactive substances from platelets constitutes an amplifying mechanism that is instrumental in immune complex deposition in acute serum sickness in rabbits [4, 51. Numerous lines of evidence indicate that PAF is released from circulating basophils. In immunized rabbits, PAF has been obtained from leukocytes in the presence of the specific antigen, o r anti-IgE antiserum [2]. The presence of histamine-containing cells has been well correlated with the release of a soluble factor [6], now known to be identical with PAF. We have presented ultrastructural evidence of rabbit platelets aggregating around degranulating basophils [2]. In addition, a factor bearing some resemblance to PAF has been obtained from a pure basophil preparation from human leukemic blood [7].
2.1 Buffers
Activators known to interact with mastocyte membrane receptors [8, 91, such as 48/80, C 5 a and neutrophil cationic proteins, were capable of releasing PAF from human leukocytes and human and rabbit tissues [lo]. Accordingly, with others [11], we suspected mastocytes as a likely source for PAF. In order to examine this hypothesis, we have studied peritoneal cells (PC) from rats and mice using various cell separation procedures. In this study, we will show that, contrary to our expectations, we have recovered PAF from macrophage-like cells and not from purified mastocytes. [I 22431
* This work was supported by INSERM: AT No. 77-79, and DGRST: ACNO.77-7-1409. Correspondence: Jacques Benveniste, INSERM U 131, 32 rue des Carnets, F-92140 Clamart, France Abbreviations: P A F Platelet-activating factor PC: Peritoneal cells HEPES: N-2-Hydroxyethylpiperazine-N-2-ethanesulfonic acid BSA. Bovine serum albumin HBCM: HEPES buffer containing calcium and magnesium ions HB: HEPES buffer SRS-A: Slow-reacting substance of anaphylaxis ECF-A: Eosinophil chemotactic factor of anaphylaxis
0Verlag Chemie, GmbH, D-6940 Weinheim, 1979
RPMI 1640 (Flow Labs., Irvine, Scotland) was prepared by dissolving 10.46 g of powder in 1 liter of distilled water and M (final) N-2-hydroxythen buffered to pH 6.8 with 5 X ethylpiperazine-N’-2-ethanesulfonicacid (HEPES) (Calbiochem, Los Angeles, CA). In some experiments, 2.5 g of bovine serum albumin (BSA) were added to RPMI (RPMI-BSA) and the p H adjusted to 7.4 with 0.1 N NaOH. A HEPES-buffered medium containing calcium and magnesium ions (HBCM) was prepared as described in [12]:for 1 liter, we used 4.77 g HEPES, 0.37 g KCl, 8.18 g NaC1, 1.00 g glucose, 0.20g CaCl, X 6H,O and 0.10 g MgCl, X 6 H z 0 . The same buffer was prepared without calcium and magnesium (HB). When 2.5 gAiter BSA were added to HBCM and HB, they were designated as HBCM-BSA and HB-BSA.
2.2 Preparation of PC One to 3-month-old Wistar rats and C57BL/6 mice of both sexes were used. PC were collected according to Bloom et al. [13]. Briefly, RPMI (10 ml for rats or 3 ml for mice) was injected into the peritoneal cavity of exsanguinated and etherkilled animals. After abdominal massage, the peritoneum was opened and the injected fluid collected with siliconized Pasteur pipettes. The cells were washed twice by centrifugation (200 X g, 5 min) and resuspended in suitable media, generally HBCM. The cells were stained by mixing 10 pl of the cell suspension with 90 p1 of 0.1% toluidine blue solution in 10% formaldehyde, SO% ethanol and 1 % acetic acid. This solution selectively stains the mastocyte granules [ 141. Differential cell counts were performed on smears stained with May-Griinwald-Giemsa solution. Histochemical staining of peroxidase-positive cells was performed as described by Preud’homme et al. [15]. In some experiments, PC were collected 48 h to 1 week after an intraperitoneal injection of distilled water [16]. Such cell preparations were devoid of mastocytes. 0014-2980/79/0S05-0409$02.S0/0
410
Eur. J . Immunol. 1979. 9: 409-415
J. M. Mencia-Huerta and J. Benveniste
2.3 Fractionation of PC 2.3.1 Ficoll gradients A stock solution of 15% (w/v) of Ficoll (Pharmacia, Uppsala, Sweden) in distilled water was buffered with 20mM HEPES and the pH adusted to 6.8 with 1N NaOH. A solution of 5% (w/v) Ficoll was obtained by addition of 2 volumes of HB to 1 volume of 15 % Ficoll solution. Gradients were prepared in 14-ml cellulose nitrate tubes (14 mm X 94 mm) with a gradient former (Isco, Lincoln, NE) using 6.5 ml of 5% and 6.5 ml of 15% Ficoll solutions. One X lo7 PC in 500 pl of RPMI were layered on the top of the gradient and centrifuged at 200 X g for 3 rnin at room temperature. Fractions of approximatively 850 pl each were collected with a fraction collector (Auto densi-flow, Buchler, Fort Lee, NJ). The fractions were then diluted 2-fold with HB-BSA, and the cells were pelleted by centrifugation at 200 X g for 15 min, washed twice with HB-BSA, sedimented by centrifugation (200 X g, 5 min) and finally resuspended in 0.4 ml HBCM-BSA or RPMI-BSA. This method was preferred to that of Sullivan et al. [17] or Orange et al. [12] which in our experiments did not yield mononuclear cell-rich fractions totally free of mastocytes.
5 min, the cells were resuspended in 200 p1 of RPMI, preheated for 10 rnin at 3 7 ° C and 1 pg/ml of ovalbumin then added. The reaction was stopped 5 min later by addition of 50 pl of the formaldehyde-containing staining solution. Cells were examined microscopically 30 min after staining and the number of degranulated vs. intact mastocytes assessed. This technique was performed either on whole PC or on Ficoll gradient-purified mastocytes.
2.6 Assay procedures PAF activity was determined in an aggregometer apparatus (Icare, Marseille, France) by aggregation of 5 X lo6 to 7 X lo6 washed rabbit platelets in 300 pl Tyrode's buffer in the presence of 5 p~ indomethacin (Sigma). It was expressed in arbitrary units as measured by the relative chart distance for the aggregation produced by 10 plof PAF-containingsolution. The preparation of platelets and the composition of buffers have been described in detail elsewhere [2]. Histamine was measured by an automatic spectrofluorometric method [ 191, and acid phosphatase with p-nitrophenyl phosphate as substrate [20] after lysis of cells with Triton X-100. Cell viability was assessed before and after each experiment by the trypan blue exclusion method.
2.3.2 Cell adherence One X lo7 cells in RPMI-BSA (2 ml) were plated in 3.5 cm diameter treated plastic petri dishes (3001 Flow) and incubated at 37 " C for 6 h. The dishes were thoroughly washed with RPMI-BSA after 2,4 and 6 h. Nonadherent cells were pooled, pelletted by centrifugation (200 X g, 5 min) and resuspended in 1 ml of RPMI-BSA.
2.4 Procedure for cell incubation The following substances were used on whole or purified PC: ovalbumin (Sigma Chemical Co., St. Louis, MO); compound 48/80 (Sigma) was dissolved in suitable media immediately before use; a stock solution of 5 mg/ml ionophore A 23187 (Elli Lilly Labs., Indianapolis, IN) was prepared in dimethyl sulfoxide and stored at - 20°C. It was diluted with saline buffer to suitable concentrations immediately before use. Equivalent dilutions of dimethyl sulfoxide were added as control. Reactions were carried out at 37 'C for 60 min, unless otherwise indicated, and were stopped by centrifugation (200 X g, 5 min) or by addition of 2.5 pl/ml of 0.2 M EDTA. In the case of plastic-adherent cells, release experiments were performed directly in petri dishes, by addition of activating substances; the incubation media were collected, centrifuged and stored. In some cases, maximal releasable PAF was obtained by overnight incubation of the cells at room temperature in HBCM-BSA at p H 10.6 [3]. All supernatants were stored at - 20 " C until assay of the released substances.
2.5 Passive sensitizationof rat PC Passive sensitization was performed according to A. ProuvostDanon et al. [18]. Briefly, 2 X l o 6 PC in 200 pl RPMI were incubated for 30 rnin at 3 7 ° C under agitation with various dilutions of IgE-rich mouse anti-ovalbumin antiserum with a titer > 1: 10000 (a gift of A. Prouvost-Danon). After 3 washings with RPMI alternated with centrifugations at 200 X g for
2.7 Characterization of PAF Aggregation of platelets by PAF was not influenced by indomethacin. In addition, two criteria to distinguish PAF from other possible aggregating agents were applied. PAF-containing solutions were incubated with phospholipase A, (Boehringer, Mannheim, FRG), as described [21], or chromatographed on silica-gel plates. To perform the thin-layer chromatography, 3 volumes of absolute ethanol wereaddedto 1volume of the PAF solution. After 25 rnin under agitation, the tubes were centrifuged at 1500 X g for 30 min. The supematants were evaporated to dryness, and PAF was recovered by addition of 200 pl of ethanol, 50 to 100 pl of which were layered on silica-gel plates 60F254 (Merck Darmstadt, FRG). The plates were developed in chloroform: methanol: water (70 :35 :7) and either stained by exposition to iodine vapor or divided into 10 X 10 mm squares. The gel of each square was recovered by scraping, and PAF activity was eluted with 100 pl of HBCM-BSA.
3 Results 3.1 Effect of the ionophore and 48/80 on unfractionated rat PC Upon addition of the ionophore A23187 or 48/80 to 5 X lo6 PC in 1 ml of HBCM-BSA, release of PAF was time-dependent and maximal after 40 min for the ionophore, 60 rnin for 48/80. The 60-rnin interval was chosen for all following exM periments. The release of PAF was abolished by 5 X EDTA or by incubation at 4 "C. By contrast with 48/80 which yielded low and variable amounts of PAF, the ionophore induced high releases from all PC populations. Cell viability was not affected, as less than 2 % of cells lost their ability to exclude trypan blue after 1 h incubation with or without ionophore or 48/80. By contrast, with PAF, histamine release was
Eur. J. Immunol. 1979. 9: 409-415
Release of platelet-activating factor from macrophages
already maximal at 5 min and at the same level with either the ionophore or 48/80 (Fig. 1).
A
We have studied the effect of increasing concentrations of ionophore on the release of PAF and histamine and mastocyte degranulation. The release of PAF was nearly maximal with 0.5 pg/ml ionophore when mastocyte degranulation was less than 14%, in contrast to histamine which closely followed degranulation (Fig. 2). Also, comparable amounts of PAF were obtained from a variety of PC showing large differences in their mastocyte numbers. Finally, PC from rats depleted of mastocytes by a previous injection of distilled water released PAF at the same level as PC from uninjected animals (Fig. 3).
%
41 1
A
A
h
2 A-
mean * s d
A A A
42.4 t 4 . 2 A
A A A
A
Units
1001
r50
Figure 3. Comparison of the release of PAF with the number of mastocytes in rat PC from different animals. (r = 0.134; not significant). PAF was obtained from 5 X 106 PC in 1 ml HBCM-BSA by incubation with 1 p g h l ionophore for 60 min and expressed as shown in Fig. 1. Mastocytes were counted microscopically afterstainingby toluidine blue. (A) PC from normal rats. ( A t )PC from rats injected with distilled water 3-7 days before experiments.
-
n
100%
al
10
20
30
40
50
60
90
T i m e (min)
Figure I . Release of histamine (---) and PAF (-) from 5 X 106rat PC in 1 ml HBCM-BSA by 1 pg/ml ionophore (A) or 1 pg/ml48/80 ( 0 ) .(0) incubation with HBCM-BSA alone, or at 4"C, or with 5x M EDTA. Histamine is expressed as the percentage of the amount found in supernatants over the total cell content (8.0 -t 3.0pg). PAF is expressed in arbitrary units, as described in Sect. 2.6. The 40-min incubation of PC with the ionophore yielded as much releasable PAF as the exposure of cells to alkaline pH. Data represent the mean f 1 standard deviation of 5 experiments.
a
r
1jBiTT-j
T u b e number
Figure 4 . Cumulative percentage of different cell types of rat PC after fractionation on 5-15 % Ficoll gradient. (A) Small lymphocytes; (B) macrophages, large lymphocytes and monocytes; (C) mastocytes; (---) concentrations of Ficoll. This figure is representative of 20 cell separation experiments. In this and all following figures, gradient fractions are numbered from top (left, abscissa) to bottom (right, abscissa).
3.2 Distribution of rat PC after Ficoll gradient separation J
I
0.05
jmmj
0.1
0.5
1
5 pglml
lonophore
Figure 2. Degranulation of mastocytes (0--0),release of PAF (A-A) and histamine (&--A) after incubation of 5 X 106 rat PC with increasing amounts of ionophore for 60 min. PAF and histamine are expressed as shown in Fig. 1. Data represent the mean k 1 standard deviation of 3 experiments (degranulation and histamine release) and of 5 experiments for release of PAF.
The distribution of PC, shown in Fig. 4, is expressed in cumulative percentages for each cell type. This method produced a good separation of mastocytes which sedimented at the bottom of the gradient, from macrophages and lymphocytes, which were more buoyant. To some extent, macrophages and large lymphocytes could be separated from small lymphocytes. Cell recovery was 70 to 80%, and the purity of mastocyte populations was between 95 and 99%. Fractions containing lymphocytes and macrophages were totally devoid of mastocytes.
412
J. M. Mencia-Huerta and J. Benveniste
Monocytes, identified as peroxidase-positive cells, accounted for 5 % of the total populations and were recovered in fractions 4 and 5 . In contrast to the results of others [22], neutrophils were very rare in our preparations (0.96 k 0.63% in 5 experiments).
3.3 Release of PAF from fractionated rat PC Exposure to alkaline p H of Ficoll fractions yielded PAF from the upper region of the gradient, corresponding to mononuclear cells, i.e. large lymphocytes, monocytes andmacrophages. PAF was associated with acid phosphatase-containing cells and was neither obtained from the top fractions corresponding to small lymphocytes nor from the lower region corresponding to mastocytes. Only the latter fractions yielded histamine (Fig. 5 ) . When fractions were incubated with 1 y g h l ionophore, PAF was again obtained only from the upper region of the gradient (Fig. 6). Cell fractions were also incubated with 1 pg/ml48/80. Neither PAF nor any other aggregating activity was detected in any region of the gradient (10 experiments), whereas release of histamine (and degranulation of masto-
Eur. J. ImmunoI. 1979. 9: 409-415
cytes) occurred as expected in the mastocyte-containing tubes (3 experiments) (Fig. 6).
3.4 Separation of mouse PC Separation of mouse PC was rather poor as compared to rats. Nevertheless, PAF coincided with acid phosphatase activity in a region where few mastocytes were present (Fig. 7, bottom). The amount of PAF and acid phosphatase activity obtained from mastocyte-depleted mice was comparable to that of normal mice (Fig. 7, top).
3.5 Effect of Ficoll fractionation on rat cell populations The following experiments were performed to check whether centrifugation in Ficoll was capable of causing cell damage. (a) The loss in cell viability, as assessed by trypan blue exclusion, was less than 2 % after the separation and washing procedures; (b) the amount of PAF released by the ionophore from unfractionated PC was identical to that of the same population, either separated and pooled back to the original amount of cells or incubated 15 min in 15% Ficoll solution and then washed; (c) mastocytes which sedimented in the 12 to 15% Ficoll fractions did not degranulate spontaneously but degranulated and released their histamine upon addition of
Figure 5. Presence of PAF (-0) (mean 4 1 standard deviation of 8 experiments), acid phosphatase (0-0) and histamine (A-A) in Ficoll gradient-fractionated rat PC. PAF was released at alkaline pH, acid phosphatase was measured after lysis of cells by Triton X-100 and histamine after boiling the cell preparations for 5 min.
Units
10-6
501
1 . 12
p%q Figure 6. Release of PAF from Ficoll gradient-fractionated rat PC or 48/80 (0); release of histamine by by 1 kg/ml ionophore (A-A) 1 pg/ml 48/80 ( C - - 0 ) . Each point represents the mean f 1 standard deviation of 6 experiments for the ionophore, 10 experiments (4WXO-PAF) and 3 experiments (48/80-histamine).
Ix
14
Tube n u m b t i
Figure 7. PAF (0-0) and acid phosphatase (0-0) from Ficoll gradient-fractionated mouse PC. Top: PC harvested 3 days after intraperitoneal injection of distilled water. Bottom: untreated mice. (W-W) total cell counts. Vertical bars represent mastocyte number. These data are representative of 5 experiments.
Eur. J . Immunol. 1979. 9: 409-415
Release of platelet-activating factor from macrophages
1 pg/ml of 48/80 (Table 1 and Fig. 6); (d) PC passively sensitized with mouse anti-ovalbumin antiserum were fractionated and incubated with 1 pg of ovalbumin. Mastocytes degranulated to the same degree, before and after passage through the gradient (Table 1). In some experiments, passive sensitizations with the anti-ovalbumin antiserum were performed after passage of mastocytes through the Ficoll gradient. Upon addition of the antigen, they degranulated as well as mastocytes from unfractionated PC (results not shown). Table 1. Effect of the separation procedure on rat peritoneal mastocytes
Hi\tamine release by 4x 80;'' Ikgrsnulation of nia\ttryte\"' Spontaneous 13) 4X. XI) O\alhuinind'
Before gradient separation
After gradient separation
63.5 t 4 9 '
66.3 t 6.0
7.7 t 3.0 70.3 t 5.0 74.0 3.3
7.1 3.9 78.9 t -1.0 73.0 f 0.8
*
*
a) Mastocytes in 200 p.l of RPMI were incubated 5 min at 37 "C with 1 pg/ml 48/80. Histamine release was assessed as described in Sect. 2.6 and expressed as percentage of total content. b) Data represent the mean t 1 standard deviation of 5 experiments. c) Counts were performed 30 min after addition of 50 p1 of the stainig solution to 200 p.l of the mastocyte suspension and degranulation expressed as percentage of degranulated vs. intact mastocytes. d) PC were sensitized by incubation for 30 min at 3 7 ° C with IgE-rich mouse anti-ovalbumin antiserum (final dilution 1 : 500). They were then exposed to 1 p g h l ovalbumin for 5 min and mastocyte degradation assessed microscopically.
3.6 Role of mastocytesin the release or inhibition of PAF Purified rat mastocytes from Ficoll gradients were pooled and incubated with 1 pg/ml of 48/80 or ionophore, or after passive sensitization with IgE-rich anti-ovalbumin antiserum, with 1 pg/ml ovalbumin. No release of PAF was obtained. To determine whether the lack of PAF release could in fact be due to the degradation of PAF or to inhibition of PAF-induced aggregation of platelets, purified mastocytes were sonicated or lysed with distilled water. The cell lysates were then incubated
4 13
with PAF (approximatively 30 units/l0 PI). No loss in PAF activity was found after 1 h at 37 "C. Also, we could not detect any inhibitory effect of the cell lysates or of supernatants from 48/80-degranulated purified mastocytes on the PAF-induced aggregation of platelets, therefore the presence of an inhibitor of platelet aggregation can be ruled out. To determine whether spontaneous release of PAF from mastocytes or its inactivation occurred during the passage of the cells through the Ficoll solution, we prepared gradients containing either 0.25% BSA as a carrier of PAF [2] or PAF (30 units/l0 pl). After deposition of PC, centrifugation and recovery of fractions, no spontaneous release of PAF or loss of activity could be demonstrated (3 experiments). 3.7 Fractionation of PC by adherence PAF was obtained from adherent rat cells by action of the ionophore or alkaline pH (Table 2); 48/80 failed to induce the release of PAF from adherent or nonadherent cells. A small amount of PAF was obtained from nonadherent cells by action of the ionophore. In an attempt to show whether PAF release occurred during the adherence of macrophage's, media from petri dishes were extracted with 3 volumes of ethanol, centrifuged, evaporated to dryness and then resuspended in 100 ~1 of HBCM-BSA. No PAF activity was recovered.
3.8 Characterization of PAF PAF obtained from macrophages before and after Ficoll gradient, migrated as a lyso-phosphatidylcholine on silica-gel thinlayer chromatography (Rf : 0.35), between sphingomyelin and phosphatidylcholine. This chromatographic behavior was identical to that of PAF from blood leukocytes (Fig. 8). Arachidonic acid, another platelet-aggregating agent, migrated with the solvent front. Three thousand units/ml of PAF from macrophages were completely inactivated, as well as PAF from leukocytes, by phospholipase A, from Naja-naja or Crotalus. PAF from macrophages or leukocytes aggregated platelets in the presence of 5 pM indomethacin, a concentration that totally suppressed an equivalent aggregation by 10 p~ arachidonic acid. +-solvent front ---------------------or
Table 2. Release of PAF from adherent and nonadherent cellsa)
.. .. H
Treatment
Adherent cells
Nonadherent cells w
Nil 3X/XO
lonophorc pH 10.0
0 Traces 35 t 9' 39 7
+
0 Traces 12 5-1 7 t6
.o
4-
Rf 0.35
. . . . . . 1
a) Rat PC were allowed to adhere on plastic petri dishes for 6 h. Nonadherent cells were recovered by washing at 2 , 4 and 6 h. 1 pg/ml 48/80 or ionophore was then added. Supernatants (1 ml) were collected after 1 h incubation at 37 "C. In some experiments, release of PAF was obtained by exposure to pH 10.6, as described [3]. b) Results are expressed in arbitrary units as the mean ? 1 standard deviation of 6 experiments.
H
2
3
4
5
6
;
m Figure 8. Diagrammatic representation of PAF on silica-gel thinlayer chromatography. (1) PAF from pig blood leukocytes; (2) PAF from rat PC before and (3) after Ficoll gradients; (4) sphingomyelin; (5) phosphatidylcholine; (6) lyso-phosphatidylcholine; (7) arachidonic acid. Solvent = chloroform :methanol : water (70 :35 : 7).
414
Eur. J. Immunol. 1979. 9: 409-415
J. M. Mencia-Huerta and J. Benveniste
4 Discussion The release of PAF was obtained by the action of the ionophore A 23187 or compound 48/80 on rat and mouse PC, a mixed population of lymphocytes, macrophages and mastocytes. The release of PAF by the ionophore was not correlated with the number of mastocytes. It was identical in mastocytedepleted PC and in whole PC population. PC were centrifuged on Ficoll gradients, and fractions containing lymphocytes and mononuclear cells (including macrophages) released PAF, but 95 % pure mastocyte preparations were inactive. PAF was present in fractions containing acid phosphatase, a marker for macrophages [23], and in adherent cells, but not in fractions containing histamine (only present in mastocytes) and in small amounts in nonadherent cell populations. The liberation of PAF by the ionophore A 23 187 was an active process which was inhibited by EDTA or low temperatures. Besides the ionophore, 48/80 or antigen also induced degranulation of purified mastocytes and the release of histamine, but failed to liberate PAF. Negative results were also observed by incubation of mastocytes at pH 10.6, a procedure which is very efficient in releasing PAF from blood leukocytes [3] or lymphomononuclear-rich PC fractions. The lack of PAF release from mastocytes could not be explained by its spontaneous release or degradation during the separation procedure. All these results exclude mastocytes and indicate that macrophages are a likely source for PAF. Besides histamine, 48/80 released PAF from whole PC - although much less than ionophore or alkaline pH - but not from fractionated PC containing either lymphocytes and macrophages, or mastocytes. This indicates that mastocytes could intervene in the release of PAF, possibly through an interaction with macrophages :degranulation of mastocytes and release of histamine (and other mediators) in a first step, then, in a second step, phagocytosis of the mastocyte granules by macrophages. This hypothesis is supported by the fact that we have obtained the release of PAF along with lysosomal enzymes, during phagocytonis of zymosan particles - another argument in favor of the macrophage origin of PAF - and observed the presence of mastocyte granules in macrophage phagosomes after incubation of whole PC with 48/80 ([24], and manuscript in preparation). A two-step process analogous to our observations could explain the well-documented IgEdependent release of slow-reacting substance of anaphylaxis (SRS-A) 1251 and eosinophil-chemotactic factor of anaphylaxis (EFC-A) [26]. Our results do not allow us to determine whether all adherent cells were involved in the release of PAF. Adherent cells are not only macrophages [27], and we cannot exclude the possibility that another cell than the macrophage could also generate PAF. These restrictions are common in all situations where cell purification could not be performed to a sufficient degree. Our macrophage-derived PAF appeared to be physicochemically identical with PAF from human, rabbit, hog and horse leukocytes, as assessed by the currently available methods of identification. The finding of a platelet-activating agent raises the question of its biochemical nature. A substance with platelet-activating activity, not further identified, was obtained from a 100% pure human basophil preparation [7]. PAF was released from human buffy coat cells with anti-IgE antiserum, C5 a or neutrophil cationic proteins, well-known activators of cells of the mastocyte-basophil series [lo], and we have confirmed (Benveniste and Camussi, unpublished results) its iden-
tity with PAF from rabbit leukocytes. Besides in blood, PAF has been described as originating from various tissues [ 10, 11, 281. PAF was not precisely identified [ l l , 281, and we could not ascertain the nature of the factor obtained from lung and kidney in the experiments reported [lo]. We know now that the finding in media from immunologically stimulated cells or organs of a platelet-activating and/or releasing substance is not a valid criterion to affirm the presence of PAF, as previously defined [21]. Other aggregating agents can be found in bathing media from lungs and kidneys (Benveniste et al., manuscript in preparation); this is the case with arachidonic acid which mimics the effect of PAF on platelets but is inhibited by indomethacin. It remains, that to date no experimental data are available to confirm the existence of different molecular species of the 1-lyso-phosphatidylcholine with platelet-aggregating activity [21]. However, until the complete molecular structure of PAF is elucidated, the identity of these PAF from leukocytes or macrophages of various origins cannot be strictly defined. Nevertheless, given the similitude of their characteristics, any existing differences should probably be minor.
Our present data are not contradictory to the well-accepted basophil origin of PAF. However, they necessitate a reconsideration of the pathophysiological role of PAF (and other mediators): far from being specific of the cells involved in the mechanisms of immediate hypersensitivity, it may be one of the numerous mediators belonging to the cells of the phagocytic group. PAF was originally described in basophils from immunized rabbits after challenge with specific antigen or anti-IgE [2]. In contrast to our present results, this release was very rapid (less than 5 min), and a factor probably identical to PAF was well correlated with histamine release 161. More evidence was provided by electron microscopical studies where platelets were shown to aggregate around degranulated basophils [2]. We have discussed the facts in favor of the release of PAF from human basophils. Our findings of the macrophage origin of PAF is reminiscent of the problem of the various origins attributed to SRS-A and ECF-A, other mediators of immediate hypersensitivity [25, 261. SRS-A was first associated with the release of histamine [29, 301 and with the IgE-mastocyte system [25]. It was later obtained from polymorphonuclear leukocytes [311 and from rat peritoneal cells, regardless of the presence of mastocytes [32]. The mononuclear cell origin of SRS-A, upon stimulation by the ionophore, has recently been ascertained [22]. In this case also, mastocytes failed to release SRS-A. By contrast, SRS-A was released from purified mastocytes upon stimulation by the ionophore A23187 [33]. ECF-A has been found in polymorphonuclear leukocytes [34]. It is therefore likely that the mediators of immediate hypersensitivity, SRS-A, ECF-A and PAF, can be released by a variety of cell types under various stimuli. Our present results add a new function to the macrophage, aggregation of platelets and release of their vasoactive amines. A cooperation between macrophages and platelets has recently been described in tumoral cell growth inhibition [35]. Yet, the different mechanisms of release and the potential pathophysiological implication of the generation of PAF by macrophages remain to be elucidated.
Received August 31,1978.
IgG-binding lymphocytes control the expression of receptors for IgM
Eur. J. Immunol. 1979. 9: 415-420
5 References 1 Siraganian, R. P. and Osler, A. G., J. Immunol. 1971. 106: 1244. 2 Benveniste, J., Henson, P. M. and Cochrane, C. G., J. Exp. Med. 1972.136: 1356. 3 Benveniste, J., Nature 1974. 249: 581. 4 Henson, P. M. and Cochrane, C. G., J. Exp. Med. 1971.133: 554. 5 Benveniste, J., Egido, J. and Gutierrez-Millet, V., Clin. Exp. Immunol. 1976. 26: 449. 6 Siraganian, R. P. andOsler, A. G., J. Immunol. 1971. 106: 1252. 7 Lewis, R. A., Goetzl, E. J., Wasserman, S. I., Valone, F. H., Rubin, R. H. and Austen, K. F., J. Immunol. 1975. 114: 87. 8 Morrison, D. C., Roser, J. F. and Cochrane, C. G., Int. Arch. Allergy Appl. Immunol. 1975. 49: 172. 9 Ranadive, N. S. and Cochrane, C. G., J. Exp. Med. 1968. 128: 605. 10 Camussi, G., Mencia-Huerta, J. M. and Benveniste, J., Immunology 1977.33: 523. 11 Kravis, T. C. and Henson, P. M., J. Immunol. 1975.115: 1677. 12 Orange,R. P.andMoore, E. G., J.lmmunol.1976.117:2191. 13 Bloom, G. D., Fredholm, B. and Maegermark, O., Acta Physiol. Scand. 1967. 71: 270. 14 Mota, I . and Dias da Silva, W., Nature 1950.186: 245. 15 Preud'homme, J. L. and Flandrin, G., J. Immunol. 1974. 113: 1650. 16 Facette, W., J. Exp. Med. 1954. I00t217. 17 Sullivan, T., Parker, K. L., Eisen, S. A. and Parker, C. W., 1. immunol. 1975.114: 1473. 18 Prouvost-Danon, A,, Wyczolkowska, J., Binaghi, R. and Abadie, A., Immunology 1975.29: 151.
Christiane SamarutO and Jean-Pierre Revillard Laboratoire d'Immunologie, INSERM U-80 and C.N.R.S. ERA 782, Lyon
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19 Ruff, F., Saindelle, A., Dutripon, E. and Parrot, J. L., Nature 1967.214: 279. 20 Linhardt, K. and Walter, K., in Bergmeyer, H.-U. (Eds.), Methods of Enzymatic Analysis, Academic Press, New York 1965, p. 776. 21 Benveniste, J., Le Couedic, J. P., Polonsky, J. and Tence, M., Nature 1977.269: 170. 22 Bach, M. K. and Brashler, J. R., J. Immunol. 1978.120: 998. 23 Cohn, Z. A. and Benson, R., J. Exp. Med. 1965.121: 153. 24 Benveniste, J. and Mencia-Huerta, J. M., Fed. Proc. 1978. 37: 1554. 25 Orange, R. P., Stechschulte, P. J. and Austen, K. F., J. Immunol. 1970.105: 1087. 26 Kay, A. B. and Austen, K. F., J. Immunol. 1971.107: 899. 27 Nathan, C. F., Asofsky, R. and Terry, W. D., J. Immunol. 1977. 118: 1612. 28 Kravis,T. C. andHenson, P. M., J. Immunol. 1977. 118: 1569. 29 Kellaway, C. H. and Trethewie, E. R., Quart. J. Exp. Physiol. 1940. 30: 121. 30 Schild, J . O., Hawkins, D. F., Mongar, J. L. and Herxheimer, J., Lancet 1951. i: 376. 31 Conroy, M. C., Orange, R. P. and Lichtenstein, L. M., J. Immunol. 1976.116: 1677. 32 Bach, M. K. and Brashler, J. R., J. Immunol. 1974.113: 2040. 33 Yecies, L. D., Wedner, H. J., Johnson, S. M. and Parker, C. W., Fed. Proc. 1978. 37: 1667. 34 Czametzki, B., Konig, W. and Lichtenstein, M. L., Nature 1975. 258: 725. 35 Johnson, R. J., Pastemack, G. R. and Shin, M. S., J. Immunol. 1977.118:494.
Human T lymphocyte receptors for IgM: control by IgG-binding lymphocytes* The capacity of human peripheral blood lymphocytes to bind antigen-IgG or antigenIgM-antibody complexes was investigated using a rosette technique with ox erythrocytes (E) coated with rabbit IgG (AG)or IgM (AM)antibodies. EAMrosette formation was achieved only in suspensions pre-incubated for 24 h at 37 "C. Addition of either EAMor EAG complexes to the culture medium was shown to prevent the formation of EAM rosettes. The inhibition was reversible, it was not due to trace IgM contaminants in the IgG antibody fraction. It was not observed when lymphocytes depleted of EAG-rosetting cells were incubated with EAG complexes. Inhibition of the expression of lymphocyte receptors for IgM can be regarded as a consequence of the modulation of surface receptors for IgG and involves an interaction between the two lymphocyte subsets bearing surface receptors for IgG and IgM, respectively. However, these experiments do not exclude the possibility that a few cells which bind EAG may lose their receptors by modulation and then express a receptor for IgM.
[I 22341
* C0ntractsD.G.R.S.T. 77.7.1381 andA.T.P. INSERM78-151. AttachCe de recherche (INSERM)
Correspondence: Jean-Pierre Revillard, HBpital E. Herriot, Pavillon P, F-69374 Lyon Cedex 2, France Abbreviations: E Ox erythrocytes EAG: Ox erythrocytes coated with rabbit IgG antibodies EA,: Ox erythrocytes coated with rabbit IgM antibodies FcyR. Lymphocyte receptor for IgG FcpR Lymphocyte receptor for IgM RFC: Rosette-forming cells TG: T lymphocytes bearing FcyR TM: T lymphocytes bearing FcpR 0 Verlag Chemie, GmbH, D-6940 Weinheim, 1979
1 Introduction Numerous cell types, including B, T and nulllymphocytes, bear readily detectable surface receptors (FcyR) for the Fc portion of aggregated o r antigen-bound IgG (reviewed in [l]).Receptors for the Fc part of native or antigen-bound IgM (FcpR) have also been detected on subsets of T [2, 31 and B cells [4, 51. FcyR and FcpR are most often expressed on different cells. Hence, two distinct subpopulations of human T cells have been defined by the presence of FcyR or FcpR and designated as TG and TM,respectively [6].These two subsets were reported to differ by several morphological [7] and 0014-2980/79/0505-0415$02.50/0
