AmericanJournal of Pathology, Vol. 138, No. 1, Janua?y 1991 Copyright © American Association ofPathologists

Cytoplasmic Lipid Bodies of Human Eosinophils Subcellular Isolation and Analysis of Arachidonate Incorporation

Peter F. Weller,* Rita A. Monahan-Earley,t Harold F. Dvorak,t and Ann M. Dvorakt From the Departments of Medicine* and Pathology,j Harvard Medical School, and the Departments of Medicine and Pathology and the Harvard Thorndike Laboratories, Charles A. Dana Research Institute, Beth Israel Hospital, Boston, Massachusetts

Lipid bodies are non-membrane-bound cytoplasmic inclusions that are prominent in leukocytes engaged in inflammatory responses. As demonstrated by electron microscopic autoradiography, lipid bodies can serve as intracellular sites of 31Harachidonic acid localization in eosinophils and other cells. To evaluate the role of lipid bodies as stores of esterified arachidonate, subcellular fractionation of lipid-body-rich human eosinophils was used to isolate lipid bodies free of other organelles. In lipid bodies isolated from 3H-arachidonate-labeled eosinophils, 3H-arachidonate was esterified almost totally in glycerolipids, predominantly in classes of phospholipids, including phosphatidylinositol and phosphatidylcholine. Lipid bodies, especially in leukocytes participating in inflammation, could represent intracellular sources of esterified arachidonate available for eicosanoid formation. (Am JPathol 1991, 138:141-148)

can be mediated by activation of protein kinase C.0 The lipids, which coalesce to form leukocyte lipid bodies in vivo, although often assumed to be triglycerides, have not been defined. In leukocytes, ultrastructural autoradiographic studies have suggested a role for lipid bodies in arachidonic acid metabolism by demonstrating that 3H-arachidonate, incorporated by eosinophils3 and neutrophils,2 was localized predominantly to intracellular lipid bodies. Similarly incorporated 3H-arachidonate was localized to lipid bodies in human alveolar macrophages and mast cells67 and in munne and guinea pig peritoneal macrophages.6 In human eosinophils and neutrophils, at times when lipid bodies were dominant sites of 3H-arachidonate localization, virtually all cell-incorporated 3H-arachidonate was esterified, primarily in phospholipids.23 Although these findings demonstrated lipid bodies to be major repositories of intracellular 3H-arachidonate and provided inferential evidence that arachidonate incorporated within lipid bodies was esterified in glycerolipids, the identities of the specific 3H-arachidonyl lipids present in leukocyte lipid bodies have not been defined. In the present study, we used methods of subcellular fractionation to isolate lipid bodies from human eosinophilic leukocytes and to evaluate the species of 3H-arachidonyl lipids present within eosinophil lipid bodies.

Experimental Procedures Materials

Lipid bodies are non-membrane-bound cytoplasmic inclusions found in many cell types,1 including neutrophils2 and eosinophils.3 Both the genesis and functions of lipid bodies, however, are largely unknown. In leukocytes, lipid body formation in vivo is a morphologic correlate of cellular participation in inflammation, as evidenced by the increased numbers of lipid bodies present in human eosinophils3 and human and rabbit neutrophils'245 engaged in inflammatory responses. Lipid body formation in neutrophils is not a manifestation of toxicity but rather

DNase (bovine pancreas, 2500 units/mg), saponin, 5bromo-4-chloroindoxyl phosphate, tetranitro blue tetraSupported in part by NIH grants A122571, A120241, and CA28834. Peter F. Weller is an Established Investigator of the American Heart Association. Accepted for publication August 28, 1990. Address reprint requests to Peter F. Weller, MD, Beth Israel Hospital, 330 Brookline Ave., Boston, MA 02215. f Weller PF, Ryeom SW, Picard ST, Ackerman SJ, Dvorak AM: Cytoplasmic lipid bodies of neutrophils: formation induced by cis-unsaturated fatty acids and mediated by protein kinase C (Submitted for publication).

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zolium (Sigma Chemical Co., St. Louis, MO); Percoll, FicollHypaque, 6% dextran in normal saline (Macrodex) (Pharmacia, Piscataway, NJ); osmium tetroxide (OS04) (Electron Microscopy Sciences, Fort Washington, PA); oil red 0 (Fisher, Medford, MA); Nile red (Eastman Kodak, Rochester, NY); cytochalasin B (Calbiochem Corp., San Diego, CA); 3H-arachidonic acid (211 Ci/mmol) (Amersham, Arlington Heights, IL); RPMI 1640, fetal calf serum (FCS), Hanks' balanced salt solution (HBSS)3 (Microbiological Associates, Walkersville, MD); and 1 -methoxyphenazine methosulfate (Research Organics, Cleveland, OH) were obtained as indicated. Sucrose buffer (5X) contained 1.35 mol/l (molar) sucrose, 100 mmol/l (millimolar) KCI, 25 mmol/I HEPES, pH 7.0.

Eosinophil Isolation and 3H-Arachidonate Labeling Eosinophils were obtained by leukophoreses from a donor with the idiopathic hypereosinophilic syndrome. After erythrocyte depletion from citrate-anticoagulated blood by dextran sedimentation, leukocytes (more than 80% eosinophils, remaining were neutrophils) were collected from Ficoll-Hypaque gradients2 and washed in Call, Mg++ free HBSS. Residual erythrocytes were lysed with hypotonic saline. For labeling, eosinophils at 2 X 1 06/ml were cultured overnight at 370C, 5% C02 in RPMI 1640, 10% FCS, with penicillin (100 units/ml), streptomycin (100,g/ml), and 4 nmol/l (nanomolar) 3H-arachidonic acid (0.9 ,uCi/ml).

Cell Disruption Sucrose-relaxation buffer, modified from solutions used for subcellular fractionation of neutrophils,89 was composed of 0.27 mol/l sucrose, 20 mmol/l KCI, 1 mmol/l Na2ATP, 1 mmol/I EGTA, 5 mmol/l HEPES, pH 7.0. Eosinophils were suspended in ice-cold sucrose-relaxation buffer at 1 to 5 X 107/ml; and cytochalasin B in dimethylsulfoxide (DMSO) was added to a final concentration of 5 gg/ml (and 0.01% DMSO). After 15 minutes at 4°C, eosinophils were subjected to nitrogen cavitation (Model 4635, Parr Instrument, Moline, IL) at 250 psi for 20 minutes at 4°C. The cavitate was collected dropwise and DNase (1 mg/ml) was added to it. Preliminary experiments indicated that disruption under these conditions yielded optimal lipid body release and recovery; the percentage of unbroken cells was usually 10% to 15% in repeated experiments. Nuclei and unbroken cells were sedimented at 800g for 5 minutes at 40C. Because buoyant lipid bodies began to rise during this centrifugation, care was taken to recover the uppermost layer and to mix uniformly the postnuclear supernatant by gentle shaking.

Subcellular Fractionation Percoll gradients, previously used to separate neutrophil microsomes and granules,8 were modified to isolate lipid bodies. Gradients were prepared by sequentially underlayering 3 ml of cavitate in sucrose-relaxation buffer (zone 11) with 3 ml of light Percoll-sucrose solution (zone ll) and 3 ml of heavy Percoll-sucrose solution (zone IV) and by overlayering with 3 ml of 8% sucrose solution (zone 1). The 8% sucrose solution (316 mOsm/kg), prepared in 0.2X calcium, magnesium-free HBSS, had a density of 1.033 g/ml. Sucrose-relaxation buffer (340 mOsm/kg), light Percoll-sucrose (14% Percoll, 20% 5X sucrose, 66% water; 395 mOsm/kg), and heavy Percoll-sucrose (68% Percoll, 20% 5X sucrose, 12% water) had densities of 1.039, 1.058, and 1.137 g/ml, respectively. All solutions were used at 40C; densities were measured at 40 to 60C with a digital density meter (Anton Parr, Fisher, Medford, MA). Gradients were centrifuged in SW-41 swinging bucket rotors (Beckman, Wakefield, MA) at 55,000g for 30 minutes at 40C. Gradient fractions of 0.75 to 1.0 ml were collected from the top by displacement with Percoll using a gradient tube fractionator (Hoeffer, San Francisco, CA). Any aggregated organelles in the pellet were not recovered.

Enzyme and Protein Assays Cytosolic lactic dehydrogenase (LDH),10 eosinophil-specific granule eosinophil peroxidase,11 eosinophil-specific and small granule arylsulfatase B,12 and alkaline phosphatase10 were assayed spectrophotometrically. Granule enzymes and alkaline phosphatase were assayed in samples after addition of Triton X-100 (final concentration, 0.1%). Any Percoll in samples were precipitated at 16,000g for 10 minutes before spectrophotometry of alkaline phosphatase and arylsulfatase assays. Protein was measured with the Bio-Rad (Richmond, CA) protein assay after precipitation of Percoll in samples as described.13 To evaluate histochemically alkaline phosphatase activity of intracellular lipid bodies, eosinophils were cytocentrifuged onto slides, fixed for 10 minutes while moist with 3.7% formaldehyde in HBSS, permeabilized with 0.05% saponin in phosphate-buffered saline, and incubated, with and without 2 mmol/l levamisole, for 30 minutes either at 370C with 0.23 mmol/l 5-bromo-4-chloro-indoxyl phosphate, 0.55 mmol/l tetranitro blue tetrazolium, 5 mmol/I sodium azide, 10 mmol/l MgCI2, and 0.15 mmol/l 1 -methoxyphenazine methosulfate in 0.1 mol/I TRIS-HCI, pH 9.514 or at room temperature with reagents (Vector red alkaline phosphatase substrate kit, Vector Laboratories, Burlingame, CA) as directed by the manufacturer. The fixation and permeabilization procedure preserved lipid bodies,

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as demonstrated by staining with 2% OS04, 0.1 mol/l imidazole, pH 7.5.15

Fluorescence and Electron Microscopy Recovery of lipid bodies during the development and application of subcellular fractionation techniques was monitored by fluorescence microscopy using the lipophilic fluorescent dye Nile red, which stains lipid bodies within cells.16'17 Isolated lipid bodies were visualized readily in coverslipped wet mounts of samples stained with an equal volume of 100 ng/ml Nile red in HBSS (diluted from a 10 ,gg/ml solution in acetone). For electron microscopy, fractions from density gradients were fixed for 0.5 to 1 hour at room temperature in 1.5% paraformaldehyde, 1.25% glutaraldehyde, 0.025% CaCI2, 0.1 mol/l sodium cacodylate buffer, pH 7.4. After centrifugation at 49,500g for 40 minutes at 40C, pellets were postfixed in 2% collidine-buffered OS04, stained en b/oc in uranyl acetate, and dehydrated and infiltrated for embedding in Epon and sectioning.2'8 Although fractions containing microsomes and granules routinely yielded pellets for examination, fractions from zone I of the gradients consistently yielded precipitated protein without identifiable lipid bodies or membranes. In many attempts to detect sedimentable lipid bodies by electron microscopy, multiple other approaches were used, including primary fixation in either 4% aqueous OS04 or phosphate buffered, ferrocyanide-reduced OS04, abbreviated dehydration and infiltration before Epon embedding, ultracentrifugation (100,000g for 24 hours) of osmium-fixed fractions, addition of carrier protein to fractions, dilution of sucrose-containing fractions to diminish their density, immobilization of pellets within specially constructed agar containers (Monahan-Earley RA, Dvorak AM, unpublished data), and examination of upper and lower aspects of pellets to avoid missing light-density pelletable material. With these varied approaches, proteinaceous materials, but neither lipid bodies nor membranes or other organelles, were sedimented repeatedly from the upper zone gradient fractions. Because Nile red staining demonstrated lipid bodies within these fractions, 200-,ul samples from pooled opalescent zone fractions were air dried overnight on previously gelatin-coated and formalin-fixed coverslips. Samples, examined by stereo-optics multiple times during drying, contained only many rounded droplets, many of which with time either shrunk and demonstrated shape changes or coalesced into larger drops. Coverslips at room temperature then were stained with oil red 0 for 10 minutes for light microscopy or were fixed for electron microscopy by exposures to 1.5% paraformaldehyde, 1.25% glutaraldehyde vapors for 24 hours, and then to

2% OS04 vapor for 48 hours. Coverslip samples, containing visible, friable osmiophilic material, were dehydrated briefly in alcohols, infiltrated in a brief propylene oxide-Epon sequence, and embedded and cured in Epon under an inverted Epon-filled BEEM capsule. En face thin sections of the embedded material were stained briefly in lead citrate and examined in a Phillips 400 electron microscope (Mahwah, NJ).

Lipid Analyses Lipids were extracted with acidified diethylether,'9 as detailed,20 except that formic acid was substituted for citric acid. Neutral lipids were resolved by thin-layer chromatography (Adsorbosil, Alltech Associates, Deerfield, IL) with petroleum ether:diethyl ether:acetic acid (90:15:1) with 3Hradioactivity quantitated with a TLC radiochromatography detector (Radiomatic, Tampa, FL). Phospholipids remained at the origin; diglycerides, free fatty acids, triglycerides, and cholesterol esters had retardation factors of 0.15, 0.53, 0.67, and 0.96, respectively. Phospholipid classes were resolved by straight-phase HPLC (Lichrosorb Si6O cation exchange column [Rainin, Woburn, MA] eluted with acetonitrile and water, as described2O), with radioactivity measured with an on-line scintillation counter (Radiomatic). Retention times for phosphatidylinositol, phosphatidylserine/phosphatidylethanolamine, phosphatidylcholine, and lysophosphatidylcholine were 1 1.5, 17.5, 30, and 34.5 minutes, respectively. Minor, unidentified phospholipids, which eluted between 32 to 40 minutes, were considered together as late-eluting phospholipids.

Results Lipid bodies have not previously been isolated from eosinophils, and no enzymatic or other protein markers associated with lipid bodies have been identified. Consequently during subcellular fractionations, lipid bodies were monitored routinely by fluorescence microscopy after staining with Nile red16'17 and selectively by transmission electron microscopy. Although lipid bodies are more buoyant than other subcellular organelles, methods of cell disruption by nitrogen cavitation and subcellular fractionation on Percoll density gradients, which resolved neutrophil granules and microsomal membranes,89 distributed lipid bodies throughout cytosolic fractions and, in addition, to lesser extents within membrane- and granule-containing fractions (data not shown). Consequently the density of the cavitate in sucrose-relaxation buffer was increased with additional sucrose, and a lesser density, isotonic 8% sucrose solution (zone 1) was included at the top of gradients to allow lipid bodies to separate from cytosolic

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Figure 1. top. Light microscopic view of oil red 0-stained, isolated eosinophil lipid bodies. Sample obtained from zone I of subcellularfractionation and air driedfor 16hours before staining with oil red O contains apurepreparation of many, variably sized, spherical, red lipid bodies (100OX). Bottom. Histochemical demonstration of alkaline phosphatase activity in intracellular eosinophil lipid bodies. Left, dark punctate intracellular lipid bodies visualized with phase-contrast microscopy. Right, identical cell showing alkaline pbosphatase reaction product in eosinophil lipid bodies, detected with indoxyl-tetranitroblue tetrazolium (135OX).

fractions. Despite these modifications, lipid bodies did not ascend freely; instead, as visualized with Nile red staining, lipid bodies in upper gradient fractions usually were clumped and clustered together amid an apparent proteinaceous matrix. Attempts to recover free, unentangled lipid bodies were not successful until we evaluated several agents that disrupt cytoskeletal elements, including cytochalasin B (microfilaments), colchicine (microtubules), and DNase I (binds actin). Treatment of cells with colchicine had no major effect, but previous treatment of cells with cytochalasin B (5 ug/ml) and addition of DNase (1 mg/ml) to the freshly collected nitrogen cavitate enabled recovery of free, buoyant lipid bodies in upper fractions. Thus lipid bodies appeared at least loosely associated with a network of cytoskeletal-associated proteins. Lipid bodies did not form a discrete band but imparted a faint opalescence to the uppermost zone. Fluorescence microscopy of Nile red-stained fractions revealed bright, punctate, -0.5- to 1-,um-sized spherical lipid bodies in all fractions of zone 1, with the greatest numbers in the uppermost fractions. Smaller numbers of lipid bodies were

found within the cytosolic and denser fractions. Membrane vesicles, visible with Nile red staining as thin annular fluorescent rings easily distinguishable from lipid bodies, were absent from fractions of zone 1, although vesicles could be visualized in fractions from other zones, especially zones Ill and IV. Electron microscopy of aldehydefixed or primary osmium-fixed fractions confirmed that fractions from zones Ill and IV contained sedimentable microsomal membranes and granules; but as noted in Methods, multiple attempts to sediment lipid bodies or other organelles from the fractions of zone I yielded only proteinaceous material with no lipid bodies and no contaminating membranes. The inherent buoyant densities of lipid bodies isolated in zone I fractions probably prevented their recovery by further centrifugation. Lipid bodies were detectable in abundance after samples from zone were air dried and fixed on coverslips. Light microscopy revealed oil red 0-stained lipid bodies of variable size (Figure 1, top). Electron microscopy demonstrated many variably dense, somewhat angular, non-membrane-bound lipid bodies (Figures 2A to C) whose morphologies had

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been observed to change during aldehyde and osmium vapor fixation and dehydration before Epon embedding. No contaminating microsomal structures were visualized by electron microscopy. Thus, by morphologic criteria, lipid bodies isolated from eosinophils were pure and free of other organelles. Biochemical marker assays (Figure 3) confirmed the absence of granule proteins from zone I fractions. Some cytosolic LDH was present within these fractions and alkaline phosphatase was present in the top two fractions containing the most lipid bodies. Because contaminating microsomal membranes repeatedly were undetectable in lipid body-containing fractions of zone 1, possible endogenous alkaline phosphatase activity of lipid bodies was evaluated histochemically on nondisrupted eosinophils. Identical localizations of alkaline phosphatase reaction products to eosinophil lipid bodies were obtained with two different detection methods, Vector red (not shown) and indoxyl-tetranitro blue tetrazolium (Figure 1, bottom), and were inhibitable with the alkaline phosphatase inhibitor, levamisole. Both morphologic and biochemical findings indicated that zone I fractions contained lipid bodies isolated without contaminating microsomal membranes or other organelles.

To evaluate the incorporation of 3H-arachidonate into lipids of eosinophil lipid bodies, eosinophils were incubated overnight with 4 nmol/l 3H-arachidonate and then subjected to subcellular fractionation. Zone I fractions containing lipid bodies were compared with fractions containing microsomal membranes. As shown in Table 1, more than one half of the 3H-arachidonate in lipid bodies was present as phospholipids, with diglycerides representing the next most abundant class of 3H-arachidonyl lipid. Little free 3H-arachidonic acid was present in lipid bodies, and triglyceride contained only a small portion of the esterified 3H-arachidonate.

Discussion Although observations of lipid bodies within neutrophils and eosinophils recorded the prominence of these nonmembrane-bound inclusions in cells associated with inflammation2-5 and documented the deposition of 3Harachidonate and other fatty acids within lipid bodies, 13 little is known about lipid body composition and function in leukocytes. Our goal was to isolate lipid bodies free of microsomal membranes to evaluate the 3H-arachidonyl

146 Weller et al AJPJanuary 1991, Vol. 138, No. 1

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cytoskeletal elements; and free, unaggregated buoyant lipid bodies could not be recovered readily from density 40 gradients unless cells were pretreated with cytochalasin 30 B before disruption and actin-binding DNAse I was added 20 after cell disruption. By electron microscopy, lipid bodies within cells often are observed encased among cytoskeletal 1 67 Second the inherent buoyant density 0~~~~ ~ ~~~~~~~~~~~~~~~~~~~~~~~ filaments.1' of isolated lipid bodies prevented their final recovery by 40 sedimentation with centrifugation. The many experimental 30 approaches that failed to sediment isolated lipid bodies 20 present in zone fractions did, however, provide evidence 10 that no sedimentable membranes or other organelles 0 contaminated the lipid bodies in zone 1. These isolated lipid bodies, after recovery on coverslips (Figure 2A to C), r-n _m Alk PhosDhatase also were free of contaminating microsomal structures 50 when examined by electron microscopy. 40 CT No proteins have been identified that can serve as 30markers of lipid bodies; and because lipid bodies can C-. 20 form within 30 minutes in stimulated leukocytes,2 de novo 10 - __ mom Km -l-it is possible that proteins associated with lipid bodies are 0 1 1111115 I I I I IiIII derived from the cytoplasm or from translocation from --I AA 'qv _ other sites. The finding of alkaline phosphatase activity in Arvisulfatase 30lipid body fractions of zone I was not anticipated. Although alkaline phosphatase was associated mostly with eosin20ophil membrane- and granule-containing fractions (Figure 103), the distribution of alkaline phosphatase (or other poI 0'-1 I I I I I I I I I I I I I I I I I I tential membrane markers) in human eosinophils has not been evaluated extensively. Two findings indicated that I;A a Peroxidase the activity detected in the most buoyant fractions was 40 not attributable to contamination with membranes. First, 30 20 as noted above, electron microscopy of lipid body-con10 taining fractions consistently failed to detect membranes, :I --"- ---m . 0 even when these fractions were processed with varied djO xUCdJJJr 4hi conditions that would have sedimented vesicles, memFraction * branes, and organelles. Second histochemical staining of Figure 3. The distribution ofprotein and enzyme markersfrom eosinophils, processed to preserve lipid bodies, directly the subcellularfractionation of human eosinophils. Results are demonstrated alkaline phosphatase activity in lipid bodies. presented as % frequency, which represents the percentage recovered in each 0. 75-mlfraction of the total recoveredfrom This in situ localization of alkaline phosphatase to eosinall gradientfractions. Zone II designates the 3 ml ofeosinophil ophil lipid bodies correlates with a previous ultrastructural cavitate in sucrose-relaxation buffer, zones I1 and IV represent the 3 ml of light and heavy sucrose-Percoll solutions, study that demonstrated alkaline phosphatase in small respectively, and zone I represents the 3 ml of overlying 8% osmiophilic intracytoplasmic, nongranule structures, which sucrose solution from which lipid bodies were isolated. appear to be lipid bodies, within eosinophils.21 Thus alkaline phosphatase activity in zone I could be attributed to lipid bodies; and alkaline phosphatase, like nonspecific lipids within lipid bodies. To isolate native lipid bodies, rather than those induced to form by incubations of leuesterase and peroxidase detected cytochemically in lipid kocytes with fatty acids2 or other agents,4 we used eobodies of other cells,' can be a lipid body-associated ensinophilic donor-derived blood eosinophils as a cellular zyme. source abundant in in vivo-formed lipid bodies. These Analyses of isolated eosinophil lipid bodies provided direct evidence that lipid bodies contain stores of esterified eosinophils contained 16 lipid bodies/cell, in excess of the lipid bodies found in normal peripheral blood neutroarachidonate. After overnight labeling with 4 nmol/l 3Hphils (-1/cell) or eosinophils (-5/cell).2 Isolation of lipid arachidonate (a concentration insufficient to induce new bodies was based on the buoyancy of these lipid-rich lipid body formation2), about one half of the 3H-arachistructures but was hindered by two attributes of lipid boddonate incorporated within lipids of lipid bodies was esies. First lipid bodies appeared to be associated with terified in specific classes of phospholipids. 3H-arachi1

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Table 1. 3H-arachidonate Incorporation Into Lipids of Isolated Eosinophil Lipid Bodies and Microsomal Membranes Lipid classes (%)

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Eosinophils (2 x 106/ml) were incubated ovemight with 4 nmol/l 3H-arachidonic acid and disrupted by nitrogen cavitation. Results of a representative experiment are presented in which 3.6 X 107 disrupted eosinophils were subjected to subcellular fractionation. Lipid body fractions included the three 1ml fractions from the top of the gradient (zone 1) and microsomal membrane fractions included the three 1 -ml fractions from the region between zones IlIl and IV. Each 1 -ml fraction was analyzed separately. Results included the total 3H radioactivity (3H-CPM) in the pooled fractions, the mean (±SEM) percentages of 3H-arachidonyl-lipids from each set of three fractions present as diglycerides (DG), triglycerides (TG), free arachidonic acid (AA), cholesterol ester (chol ester), and phospholipids (PL), and the mean (±SEM) percentages of 3H-arachidonyl phospholipids present as phosphatidylinositol (PI), phosphatidylcholine (PC), phosphatidylserine and phosphatidylethanolamine (PS/PE) and minor, late-eluting phospholipids (Late).

donate was distributed approximately equally among the inositol, choline, and ethanolamine/serine classes of phospholipids in lipid bodies in a similar, but not identical, pattern to that found in membranes. That eosinophil lipid bodies are rich in phospholipids has been suggested by observations with Nile red. With fluorescein filters, Nile red fluoresces yellow-gold in the presence of neutral lipids; with rhodamine filters it fluoresces red in the presence of phospholipids.17 Nile red-stained eosinophil lipid bodies, both intracellularly and when isolated, were most intensely fluorescent when viewed with rhodamine filters. Because not all lipid bodies were recovered within zone of the density gradients and because some disruption or solubilization of non-membrane-bound lipid bodies during isolation cannot be excluded, quantitation of the arachidonate present in the isolated zone lipid bodies underestimates the amount of arachidonate incorporated within these structures. These studies with isolated lipid bodies, therefore, do not provide a determination of the relative amounts of arachidonate present in lipid bodies in comparison with membranes but do establish that lipid bodies represent a nonmembraneous intracellular reservoir of esterified arachidonate. The presence of arachidonate esterified in phospholipids, including phosphatidylcholine and phosphatidylinositol, within lipid bodies of eosinophils would provide a localized, intracytoplasmic pool from which arachidonate could be mobilized for conversion to eicosanoids. In both eosinophils and neutrophils, lipid bodies are more prominent when these cells are engaged in or associated with inflammatory responses.25 Leukocytes, in sites of inflammation, are subject to activating cellular ligands and lipids and are likely to be involved in stimulated cellular responses, including eicosanoid synthesis and release. Findings with neutrophils have indicated that lipid body formation represents a specific cellular response mediated by protein kinase C activation.4 In neutrophils, several activators of protein kinase C (cis fatty acids, diglycerdes, and phorbol esters) induced lipid body formation, and inhibitors of protein kinase C blocked lipid body formation induced by these protein kinase C activators. Furthermore

lipid bodies can be sites of localization of eicosanoidforming enzymes, as evidenced by the immunochemical and biochemical localization of prostaglandin H synthase (cyclooxygenase) to lipid bodies in 3T3 fibroblasts (Weller PF, unpublished observations). Our present demonstration that isolated eosinophil lipid bodies contain 3H-arachidonate within their phospholipids, in conjunction with our previous autoradiographic demonstration that lipid bodies in eosinophils,3 like those in neutrophils2 and other cells,1'6'7 can be a predominant site of localization of incorporated 3H-arachidonate, indicate that lipid bodies may constitute a distinct, nonmembraneous pool of esterified arachidonate from which incorporated arachidonate can be mobilized for eicosanoid formation.

Acknowledgments The authors thank Sandra W. Ryeom, Susanne Picard, Patricia Estrella, Linda LeTourneau, Kathryn Pyne, and V. Susan Harvey for their technical assistance.

References 1. Galli SJ, Dvorak AM, Peters SP, Schulman ES, MacGlashan Jr. DW, Isomura T, Pyne K, Harvey VS, Hammel I, Lichtenstein LM, Dvorak HF. Lipid bodies. Widely distributed cytoplasmic structures that represent preferential nonmembrane repositories of exogenous [3H]arachidonic acid incorporated by mast cells, macrophages, and other cell types. In Prostaglandins, Leukotrienes, and Lipoxins. Bailey JM, ed. New York, Plenum Press, 1985, pp 221-239 2. Weller PF, Ackerman SJ, Nicholson-Weller A, Dvorak AM: Cytoplasmic lipid bodies of human neutrophilic leukocytes. Am J Pathol 1989, 135:947-959 3. Weller PF, Dvorak AM: Arachidonic acid incorporation by cytoplasmic lipid bodies of human eosinophils. Blood 1985,

65:1269-1274 4. Coimbra A, Lopes-Vaz A: The presence of lipid droplets and the absence of stable sudanophilia in osmium-fixed human leukocytes. J Histochem Cytochem 1971, 19:551-557

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