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Am J Physiol Heart Circ Physiol 278: H269-H276, 2000;
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Vol. 278, Issue 1, H269-H276, January 2000

Endothelial cells potentiate phagocytic killing by macrophages via platelet-activating factor release

Tetsuhiro Owaki, Avedis Meneshian, Kosei Maemura, Sonshin Takao, Dajie Wang, Katherine C. Fuh, Gregory B. Bulkley, and Andrew S. Klein

Department of Surgery, Johns Hopkins University School of Medicine, Baltimore, Maryland 21287-4685


    ABSTRACT
TOP
ABSTRACT
INTRODUCTION
MATERIALS AND METHODS
RESULTS
DISCUSSION
REFERENCES

The immunomodulatory function of endothelial cells (EC) includes the initiation of leukocyte margination, diapedesis, and activation through the upregulation of various cell surface-associated molecules. However, the effect that EC have on the phagocytic function of neighboring monocytes and macrophages is less well described. To address this issue, microvascular EC were cocultured with murine peritoneal macrophages, first in direct contact, then in a noncontact coculture system, and macrophage phagocytosis and phagocytic killing were assessed. The presence of increasing concentrations of EC resulted in a dose-dependent increase in macrophage phagocytic killing. This stimulatory effect was inhibited in a dose-dependent manner by the pretreatment of macrophage/EC cocultures with WEB-2086 or CV-6209, specific platelet-activating factor (PAF)-receptor antagonists, but not by anti-tumor necrosis factor-alpha , anti-interleukin (IL)-1alpha , or anti-IL-1beta . Furthermore, the effect was reproduced in the absence of EC by the exogenous administration of nanomolar concentrations of PAF. Microvascular EC potentiate macrophage phagocytic killing via the release of a soluble signal; PAF appears to be an important component of that signal.

phagocytosis; endothelium; immunomodulation; cytokines


    INTRODUCTION
TOP
ABSTRACT
INTRODUCTION
MATERIALS AND METHODS
RESULTS
DISCUSSION
REFERENCES

THE INGESTION AND DESTRUCTION of circulating pathogens by bone marrow-derived phagocytes is essential to the survival of higher eukaryotes. Although certain components of the immune response against foreign organisms are quite antigen specific, the effector mechanisms employed by phagocytes to destroy foreign targets are generally not antigen specific and entail the disruption of membranes, proteins, and DNA (4), usually within phagosomes. This lack of effector specificity necessitates the evolution of regulatory mechanisms that can selectively stimulate or suppress the immune response, thereby limiting host tissue injury from aberrant local and systemic inflammatory processes.

Endothelial cells (EC) modulate immune function in a variety of ways; through the orderly expression of cell surface-associated molecules such as P-selectin (12, 15), intercellular adhesion molecule 1 (30, 38), and platelet-activating factor (PAF) (19), they initiate neutrophil rolling, adhesion, diapedesis, and activation in response to tissue injury and infection. A number of EC-secreted soluble mediators, such as interleukin (IL)-1 (17, 18, 22), IL-6 (33), colony-stimulating factors (3, 20, 31, 32), and PAF (5, 7), have also been found to modulate various components of the immune response. Moreover, we have recently reported that hepatic sinusoidal EC, when cocultured with neighboring Kupffer cells (KC), can potentiate KC phagocytosis and phagocytic killing (28). However, the purity of the primarily harvested cells used in this study, the generalizability of our findings to extrahepatic EC and macrophages, and, most importantly, the mechanism of this stimulatory response remain to be elucidated. This study was undertaken therefore to determine whether microvascular EC, when cocultured with macrophages, could affect murine peritoneal macrophage phagocytosis and/or phagocytic killing and to identify the potential mediator(s) of this effect.


    MATERIALS AND METHODS
TOP
ABSTRACT
INTRODUCTION
MATERIALS AND METHODS
RESULTS
DISCUSSION
REFERENCES

Macrophages. Cells from a murine peritoneal macrophage cell line [IC-21; American Type Culture Collection (ATCC), Rockville, MD] were grown in a 37°C, 5% CO2-95% air incubator in 75-cm2 tissue culture flasks containing 10 ml of RPMI 1640 (GIBCO, Grand Island, NY) with 10% fetal bovine serum (FBS), 25 mM HEPES, 250 U/ml penicillin, and 250 µg/ml streptomycin. At the time of cell harvest, flasks were washed twice with Ca2+/Mg2+-free phosphate-buffered saline (PBS) to remove nonadherent, nonviable cells. This same buffer (10 ml) was then added to each flask and incubated for an additional 10 min at 37°C. Gentle shear force was then applied by lightly tapping the sides of each flask until most of the cells were detached into suspension, as viewed by phase-contrast microscopy. This cell suspension was then collected and centrifuged at 1,100 rpm for 10 min, and the pellet was resuspended in 2 ml of RPMI 1640. A 100-µl aliquot of this suspension was then diluted into 900 µl of a 0.4% solution of trypan blue stain (GIBCO), and viable (nonstained) cells were counted on a hemacytometer (viability >98%). Appropriate dilutions of the stock suspension were then prepared to obtain a final cell concentration of 4 × 105 cells/ml. A 1-ml aliquot of this suspension was then added to each of six 3.0-cm wells on a Multiwell tissue culture plate (Becton Dickinson Labware, Franklin Lakes, NJ) and incubated at 37°C for 2 h.

Endothelial cells. EC from a murine microvascular EC line (SVEC, ATCC no. CRL-2181; generously provided by Dr. Kathryn O'Connell, Johns Hopkins Medical Institutions, Baltimore, MD) were chosen for this experiment because they are well characterized (25) and, more importantly, are derived from the microvasculature (of mouse lymph nodes) and therefore are perhaps more representative of the microvascular EC participating in the early trigger of this particular immune response in vivo. SVEC were grown in a 37°C, 5% CO2-95% air incubator in 75-cm2 tissue culture flasks containing 10 ml of RPMI 1640 with 10% FBS, 25 mM HEPES, 250 U/ml penicillin, and 250 µg/ml streptomycin.

Primarily harvested, large-vessel EC derived from human umbilical vein (HUVEC) were also used in a number of the experiments to determine whether our findings were unique to the SVEC line or more generally applicable. HUVEC were harvested from human umbilical cords by collagenase treatment using an adaptation of a previously described method (11). All harvesting procedures were performed using a sterile technique. Briefly, the outer surface of fresh human umbilical cord sections (1-3 h postpartum) was washed with Ca2+/Mg+-free PBS. The hub of a 60-ml syringe filled with this same buffer was then inserted into one end of the umbilical vein, and 60 ml of PBS were infused to wash out clotted blood. The end opposite the syringe was then occluded, and 60 ml of a 0.1% solution of collagenase type II (Worthington Biochemical, Lakewood, NJ) dissolved in Ca2+/Mg+-free PBS were gently infused. The syringe was then removed and the open end also clamped. The umbilical cord was then placed in a beaker filled with 500 ml of warm PBS and incubated in a 37°C water bath for 20 min. After the first 10 min of incubation, the cord was gently massaged and replaced into the PBS bath. After 20 min of incubation, the clamps were removed, and the collagenase cell suspension was allowed to drain slowly into a 50-ml tube. This tube was then centrifuged for 10 min at 1,000 rpm, and the pellet was resuspended in 10 ml of a stock EC growth medium [EGM, Clonetics, San Diego, CA; medium was prepared as directed by the manufacturer, except that penicillin (250 U/ml) and streptomycin (250 µg/ml) were used instead of gentamicin and amphotericin B]. This cell suspension was then transferred to a 75-cm2 tissue culture dish that had been previously coated with 25 µg of human plasma fibronectin (GIBCO) dissolved in PBS, and it was incubated at 37°C in 5% CO2-95% air.

At the time of cell harvest, all EC-filled flasks were washed twice with Ca2+/Mg2+-free PBS to remove nonadherent cells. A 0.05% trypsin/EDTA solution (5 ml) was used in combination with gentle shear force (tapping) to bring the remaining cells into suspension, and 5 ml of FBS were then added to terminate the trypsin degradation. This suspension was then centrifuged at 1,100 rpm for 10 min, and the pellet was resuspended in 2 ml of RPMI 1640 with 10% FBS (or EGM for the HUVEC). A 100-µl aliquot of this suspension was then diluted into 900 µl of a 0.4% solution of trypan blue, and cells were counted on a hemacytometer (viability >98%). Appropriate dilutions were then prepared to achieve the required concentration (2 × 105-2 × 106 cells/ml) for each day's experiment.

A fibroblast cell line (DCEK; generously provided by Dr. Drew Pardoll, Johns Hopkins Medical Institutions) was used as a negative control for the EC component of the coculture system. Fibroblasts were incubated and harvested in a manner identical to that for the SVEC line.

Yeast cell targets. Candida parapsilosis were maintained aerobically on Sabouroud agar plates. An aliquot of this yeast was used to inoculate a 30-ml solution of trypticase soy broth, which was then incubated aerobically at 37°C for 18 h. After incubation, the yeast cells were washed twice with Gey's balanced salt solution and centrifuged at 3,100 rpm for 10 min. The pellet was then resuspended in 2 ml of RPMI 1640 with 10% FBS and 25 mM HEPES. Viable cells were counted on a hemacytometer using a 0.4% solution of trypan blue (viability >95%), and appropriate dilutions of the stock cell suspension were prepared to achieve a final concentration of 8 × 107 yeast cells/ml.

Contact coculture system. After 2 h of incubation of the macrophages in the 3.0-cm tissue culture wells, existing media were replaced with 1 ml of fresh medium. A 1-ml aliquot of the EC suspension (or a control solution of either RPMI 1640 with 10% FBS and 25 mM HEPES for SVEC/fibroblast experiments or EGM for HUVEC experiments) was then added directly to each macrophage well, and the two cell lines were coincubated in direct contact for 24 h at 37°C. The previously prepared suspension of C. parapsilosis (50 µl; 8 × 107 cells/ml) was then added to provide a yeast-to-macrophage ratio of 10:1, and this preparation was incubated for 60 min at 37°C in 5% CO2-95% air.

Noncontact coculture system. After 2 h of incubation of the macrophages in the 3.0-cm tissue culture wells, the existing media were replaced with 1 ml of fresh medium. Cyclopore cell culture inserts (0.4 µm; Becton Dickinson Labware) were then placed into these 3.0-cm wells and filled with 1 ml of EC suspension (or a control solution, same as in Contact coculture system), thereby allowing the two cell lines to share medium without direct cell-cell contact. After 24 h of coincubation at 37°C, the EC-filled inserts were removed and 50 µl of the yeast suspension (8 × 107 cells/ml) were added to the macrophage wells, thereby providing a 10:1 ratio of yeast to macrophages. This preparation was then incubated for 60 min.

Assessment of phagocytosis and phagocytic killing of C. parapsilosis by macrophages using the acridine orange-crystal violet staining assay. After 60 min of coincubation of yeast and macrophages, the media were removed and the tissue culture wells were gently rinsed with Gey's solution (pH 7.3) and stained by the method of Pruzanski and Saito (29) for 45 s with 0.01% acridine orange (Fisher Scientific, Pittsburgh, PA) dissolved in Gey's solution. The wells were then rinsed twice with Gey's solution and stained for an additional 30 s with 0.05% crystal violet (Sigma Chemical, St. Louis, MO) dissolved in 0.15 M NaCl. Finally, the wells were washed twice more with Gey's solution and this wet preparation covered with a plastic coverslip for microscopy. Macrophages were scanned under the fluorescence microscope (Carl Zeiss) with fluorescence filters (excitation 485 nm, dichroic 510 nm). Under these conditions, viable intracellular C. parapsilosis cells appear green, whereas dead yeast cells appear bright red/orange (Fig. 1) (29, 34). The correlation between fluorescence and viability (29, 34) was again confirmed by standard curves prepared by mixing varying proportions of live and dead (boiled for 30 min) organisms and validated by subsequent quantitative culture (data not shown). (Boiling for 30 min consistently yielded a population of 100% dead cells that were nonetheless recognizable morphologically and therefore could be counted microscopically.) In all experiments, except the initial experiments involving direct-contact coculture, in which it would have been impossible, the microscopist was blinded with respect to the treatment group at the time of the fluorescence analysis.


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  		Fig. 1.   Fluorescence micrograph depicting acridine orange-crystal violet assay for macrophage (Mphi ) phagocytosis and phagocytic killing of Candida parapsilosis. Note intracellular dead (orange; large arrows) and live (green; small arrow) yeast.

Phagocytic activity was expressed as the percentage of macrophages containing one or more ingested Candida organisms, regardless of whether the yeast were alive or dead (200-300 macrophages were assayed per tissue culture well). Intracellular candidacidal activity (ICCA) denotes the percentage of the total number of intracellular Candida that had been killed (i.e., stained orange rather than green; 200-300 yeast were assayed per tissue culture well). In the initial direct-contact coculture experiments, the total number of macrophages in each well could not be accurately counted because of difficulty in discriminating them from adjacent SVEC. Therefore, only ICCA was assessed in these early experiments.

Measurement of tumor necrosis factor-alpha production. Tumor necrosis factor (TNF)-alpha levels in the cell culture supernatants were measured using a standard cytotoxic assay against a TNF-alpha -sensitive mouse fibroblast cell line (WEHI 164; ATCC) as previously described (9). Briefly, 5 × 104 cells of WEHI 164 in 100 µl of RPMI with 10% FBS were placed in each of 96 wells on a flat-bottom plate and pretreated for 2 h with actinomycin D (0.5 µg/ml; Sigma Chemical) at 37°C. During this incubation period, solutions of recombinant murine TNF-alpha (R&D Systems, Minneapolis, MN) were prepared at various concentrations (1-1,000 pg/ml) to obtain standard curves for each experimental trial. After this 2-h incubation, existing media were removed and 100 µl of the coculture supernatant samples (obtained at the end of a 24-h coculture period) or the previously prepared TNF-alpha standards were added to each of the 96 wells. Each well was repeated in triplicate. This preparation was then incubated for an additional 18 h. After this period, 20 µl of 3-(4,5-dimethylthiazol-2-yl)-2,5-diphenyltetrazolium bromide (Sigma Chemical) at a concentration of 2 mg/ml in Hanks' balanced salt solution was added to each well and incubated for an additional 4 h. The medium was then removed, and 200 µl of DMSO were added to each well to dissolve the purple formazan crystals. Fifteen minutes later, 100 µl of the supernatant were transferred to each of 96 corresponding wells on a fresh flat-bottom plate and the absorbance of each well was determined at 550 nm on a microplate reader (Molecular Devices, THERMOmax, Menlo Park, CA).

Experimental protocols. In all experiments, macrophages were provided yeast for 60 min, on the basis of previously described dose- and time-response curves (34), and phagocytosis and killing were then measured. The first experiments assessed control values for phagocytosis and killing by macrophages alone. The second group of experiments was conducted to determine whether contact coculture of these macrophages with SVEC could alter macrophage phagocytosis or killing. Finally, these coculture experiments were conducted using the noncontact coculture system. After the significant increase in macrophage phagocytic killing on coculture with SVEC was noted, the generalizability of these results was assessed by repeating noncontact coculture experiments with the following additional combinations of cells: HUVEC plus macrophages and fibroblasts (negative controls) plus macrophages.

We then evaluated the potential role of PAF as a mediator of the stimulatory effect noted with noncontact coculture of macrophages and SVEC, because the generation of PAF by EC is well documented (5, 7). Macrophages alone and SVEC/macrophage noncontact cocultures were pretreated with varying doses of WEB-2086 (0-20 µM; Boehringer Ingelheim Pharmaceuticals, Ridgefield, CT), and CV-6209 (0-1 µM; Biomol, Plymouth Meeting, PA), two highly specific PAF-receptor antagonists, for 24 h before yeast challenge. In a separate series of experiments, sets of macrophage plates (without SVEC) were pretreated with varying doses of PAF (0.2-20 nM; Calbiochem, San Diego, CA) for 24 h before yeast challenge in an attempt to replicate the effect of SVEC coculture.

We also examined the potential role of TNF-alpha as a mediator of this stimulatory response, because this mediator has been shown to enhance phagocytosis and killing in human macrophages (1, 13, 37), is also known to be released by activated macrophages (35), and could conceivably be acting in this system as a positive feedback stimulant of EC PAF generation (6, 8). One group of macrophages and one of the SVEC/macrophages in noncontact coculture were pretreated with murine anti-TNF-alpha (1 µg/ml; R&D Systems) 24 h before yeast challenge and the assessment of phagocytosis and killing. Another set of macrophages was pretreated with PAF (20 nM) for 24 h in the presence and absence of anti-TNF-alpha (1 µg/ml). Finally, dishes of macrophages (without SVEC) were exposed to varying concentrations of recombinant murine TNF-alpha (5-500 units; R&D Systems) for 6 h before yeast challenge and the assessment of phagocytosis and killing. In a separate series of experiments, TNF-alpha production and release by macrophages was measured in a control group of macrophages alone and in a group consisting of the SVEC/macrophage noncontact cocultures at various ratios (1:1 to 5:1). After a stimulatory effect of PAF on phagocytic killing had been observed, we assayed TNF-alpha production by macrophages after 24 h of pretreatment with various doses of PAF (0.2-20 nM) in the absence of EC and also assayed the production of TNF-alpha over a 24-h time course in response to a 20 nM dose of PAF.

The potential roles of IL-1-alpha and IL-1-beta as mediators of this stimulatory response were also screened by pretreating macrophages alone and SVEC/macrophage noncontact cocultures with murine anti-IL-1alpha or anti-IL-1beta (1 µg/ml; R&D Systems) for 24 h before yeast challenge. Phagocytosis and killing were then assessed.

Statistical analysis. Data represent 200-300 cells assayed per tissue culture well on each of three separate experiment days and are expressed as means ± SD of the percentage of control values. Apparent individual differences between discrete treatments were assessed for statistical significance by a paired t-test. Dose-response curves were evaluated by one-way ANOVA. Values of P < 0.05 were considered significant.


    RESULTS
TOP
ABSTRACT
INTRODUCTION
MATERIALS AND METHODS
RESULTS
DISCUSSION
REFERENCES

EC coculture potentiated macrophage phagocytic killing in a dose-dependent manner. Contact coculture of SVEC with macrophages appeared to stimulate ICCA at all cell ratios, increasing in a dose-response relationship from the 1:1 to 5:1 ratios of SVEC to macrophages (Fig. 2A), although this clear trend did not reach statistical significance. Noncontact coculture of SVEC with macrophages had little effect on macrophage phagocytic activity. However, like direct coculture, it did stimulate ICCA at all cell ratios, increasing in a similar dose-response manner from the 1:1 to 5:1 ratios of SVEC to macrophages (Fig. 2B), and did reach statistical significance. On the other hand, noncontact coculture of HUVEC plus macrophages and fibroblasts plus macrophages did not affect phagocytosis or phagocytic killing (Fig. 2, C and D).


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  		Fig. 2.   Macrophage phagocytosis (Mphi ) and intracellular candidacidal activity (ICCA) after direct coculture with murine vascular endothelial cells (SVEC) (A) and after noncontact coculture with SVEC (B), human umbilical vein endothelial cells (HUVEC) (C), and fibroblast cells (DCEK) (D). Each point represents 200-300 cells assayed on each of 3 separate experiment days. Data are expressed as means ± SD of %control for the 3 experiments. * P < 0.05 vs. Mphi alone (1-way ANOVA). Mean absolute values (±SD) for phagocytosis and ICCA in control macrophages for this series of experiments were 45.5 ± 6.0% and 28.5 ± 8.0%, respectively.

Pretreatment of macrophages with WEB-2086 or CV-6209 dose dependently blocked the stimulation of macrophage phagocytic killing provided by SVEC coculture. Whereas endogenous PAF inhibition by pretreatment of SVEC/macrophage cocultures with WEB-2086 or CV-6209, two specific PAF-receptor antagonists, did not affect phagocytosis (Fig. 3A), it did dose dependently ablate all of the increase in ICCA seen in response to coculture with SVEC (Figs. 3B and 4). Moreover, like coculture with SVEC, the exogenous administration of PAF to macrophages alone (in the absence of SVEC) before yeast challenge did not affect macrophage phagocytic activity but did cause significant, dose-dependent increases in phagocytic killing (Fig. 5). A 20 nM pretreat-ment dose of PAF stimulated ICCA to nearly the same degree as the SVEC/macrophage (1:1) noncontact coculture.


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  		Fig. 3.   Macrophage phagocytosis (A) and ICCA (B) in Mphi alone and Mphi  + SVEC noncontact cocultures after pretreatment with WEB-2086 [20 µM; a specific platelet-activating factor (PAF)-receptor antagonist], anti-tumor necrosis factor-alpha (anti-TNF-alpha ; 1 µg/ml), anti-interleukin-1alpha (anti-IL-1alpha ; 1 µg/ml), and anti-interleukin-1beta (anti-IL-1beta ; 1 µg/ml). WEB-2086 completely abolished the increase in macrophage phagocytic killing seen after coculture with SVEC. Each bar represents 200-300 cells assayed on each of 3 separate experiment days. Data are represented as means ± SE of %control for the 3 experiments. * P < 0.05 vs. Mphi alone (paired t-test); +P < 0.05 vs. Mphi  + SVEC control (paired t-test).



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  		Fig. 4.   ICCA of Mphi alone and Mphi  + SVEC cocultures after pretreatment with varying doses of WEB-2086 (0-20 µM) (A) and CV-6209 (0-1 µM) (B), 2 specific PAF-receptor antagonists. Each point represents 200-300 cells assayed on each of 3 separate experiment days. Data are expressed as means ± SD of %control for the 3 experiments. * P < 0.05 vs. Mphi  + SVEC without antagonist (1-way ANOVA).



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  		Fig. 5.   Macrophage phagocytosis and ICCA after pretreatment with varying doses of PAF. Each point represents 200-300 cells assayed on each of 3 separate experiment days. Data are expressed as means ± SD of %control for the 3 experiments. * P < 0.05 vs. control Mphi alone (paired t-test). Mean absolute values (±SD) for phagocytosis and ICCA in control macrophages for this series of experiments were 66.3 ± 23.3% and 25.1 ± 10.3%, respectively.

TNF-alpha did not affect macrophage phagocytosis or phagocytic killing. Pretreatment of macrophage or SVEC/macrophage noncontact cocultures with anti-TNF-alpha did not alter phagocytic function (Fig. 3A) or ICCA (Fig. 3B) in either group. Specifically, anti-TNF-alpha did not inhibit the stimulation of macrophage phagocytic killing noted on coculture with SVEC. Moreover, the presence of anti-TNF-alpha did not alter the phagocytic activity (Fig. 6A) or ICCA (Fig. 6B) of PAF-pretreated macrophages. Furthermore, the exogenous administration of varying doses of TNF-alpha had no impact on macrophage phagocytosis or killing (Fig. 7). TNF-alpha production and release by untreated macrophages (0.31 ± 0.21 U/ml) was minimal (but still greater than the 2 × 10-3 U/ml sensitivity threshold for this assay) and did not increase significantly with SVEC noncontact coculture. Moreover, pretreatment of macrophages with PAF did not alter TNF-alpha production, regardless of the dose of PAF or the point in time along the 24-h course (standard curve and additional data not shown).


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  		Fig. 6.   Effect of anti-TNF-alpha on phagocytosis (A) and ICCA (B) of PAF-pretreated macrophages. No significant effect on phagocytosis or killing is noted after administration of anti-TNF-alpha . Each bar represents 200-300 cells assayed on each of 3 separate experiment days. Data are expressed as means ± SD of %control for the 3 experiments. * P < 0.05 vs. Mphi alone (by paired t-test).



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  		Fig. 7.   Macrophage phagocytosis and ICCA after pretreatment with TNF-alpha . No significant differences in phagocytosis or killing were noted. Each point represents 200-300 cells assayed on each of 3 separate experiment days. Data are expressed as means ± SD of %control for the 3 experiments.

Anti-IL-1alpha and anti-IL-1beta did not affect macrophage phagocytosis or phagocytic killing in our model. Pretreatment of macrophage and SVEC/macrophage noncontact cocultures with either anti-IL-1-alpha or anti-IL-1beta did not alter phagocytosis or killing (Fig. 3, A and B).


    DISCUSSION
TOP
ABSTRACT
INTRODUCTION
MATERIALS AND METHODS
RESULTS
DISCUSSION
REFERENCES

We have found that coculture of SVEC and macrophages does not influence macrophage phagocytic activity but does increase phagocytic killing in a dose-dependent manner. The finding that EC and immune cells should interact in this fashion in vitro is not surprising, because EC comprise the interface at which immune cells and infection/injury meet. Such interactions are not unprecedented; we have previously reported (28) that the direct-contact coculture of primarily harvested, hepatic sinusoidal EC with neighboring KC results in increased KC phagocytosis and phagocytic killing efficiency and that this stimulatory effect is not oxidant mediated (28). Nonetheless, the purity of these primarily harvested cells, the generalizability of our findings to extrahepatic EC and macrophages, and, most importantly, the mechanism of this stimulatory response had yet to be elucidated. Primarily harvested cells, such as the hepatic sinusoidal EC and KC used in the aforementioned study, can never be composed of purely one cell type but instead are a mixture of different cells in varying proportions. For example, hepatic EC and KC are obtained by the homogenization of whole liver and subsequent separation on the basis of their differential adhesion to tissue culture flasks. This process results in a KC population that is >90% pure (by morphology) and a hepatic EC-enriched cellular fraction composed primarily of hepatic sinusoidal EC (60-80%) but also of fibroblasts and stellate cells (20-40%). In such preparations, it is impossible to ensure that it is the EC alone that mediate this stimulatory effect of coculture. To ensure the purity of the cells in our present study, we chose two well-characterized cell lines. The murine peritoneal macrophage cell line IC-21 shares many of the characteristics of normal peritoneal macrophages (21, 23) and offers the additional advantage over primary cell culture of ensuring cell purity and avoiding the premature activation of macrophages necessitated by the harvesting of primary cells. The SVEC EC line (25), derived from the microvasculature of mouse lymph nodes, shares many characteristics with normal vascular EC, including their morphology, cell-surface associated markers, nutritional requirements, secretory properties, and response to growth factors (25), thus providing a functionally relevant model. Because the majority of EC-immune cell interactions normally occur in the postcapillary venules, and not along the walls of larger vessels (from which primary endothelial cell cultures are generally obtained), the use of microvascular EC here is relevant. Furthermore, the fact that these SVEC come from lymph nodes suggests the possibility that these cells might be more representative of the cells participating in the early microvascular immune response in vivo.

The second issue we wanted to address was whether the findings obtained with primarily harvested hepatic EC and KC were applicable to EC and macrophages in general. Because the liver is, quantitatively, the most important site of reticuloendothelial system function, accounting for 80% of the clearance and killing of bacteria circulating systemically (14), it would not be unreasonable to suspect that this stimulatory interaction might be limited to this unique environment. By using peritoneal macrophages and lymph node-derived microvascular EC lines in this study, we were able to evaluate the generalizability of these results. The fact that large vessel-derived EC (HUVEC) did not reproduce this response supports the notion that this process is specific to a subpopulation of specially differentiated EC that line the microvasculature (including the liver sinusoids), where much of the immune response is thought to occur. Certainly, the human umbilical vein wall has little known immune function.

The noncontact coculture system provided a controlled environment in which to study one component of the EC-macrophage interaction. This noncontact system, compared with direct coculture or in situ preparations, eliminates the confounding variables of direct cell-cell, cell-basement membrane, and cell-extracellular matrix interactions (2, 16), thereby excluding those interactions not mediated by diffusible factors. A major disadvantage of this approach is that elimination of these other variables precludes the evaluation of their roles in this response.

The increase in phagocytic killing efficiency seen after noncontact coculture of macrophages with SVEC was inhibited in a dose-dependent manner by WEB-2086 or CV-6209, two specific PAF-receptor antagonists, and was reproduced, dose dependently, in the absence of SVEC, by the exogenous administration of PAF in nanomolar concentrations. These findings suggest that PAF is the soluble signal that mediates this stimulatory response. In previous studies, PAF has been reported to dose dependently enhance the phagocytosis of FITC-labeled beads by murine peritoneal macrophages (10). (Interestingly, we saw no effect of PAF on phagocytic uptake.) However, we are unaware of prior studies of the effects of PAF on phagocytic killing. The use of the acridine orange-crystal violet assay (29, 34) allows the quantitative discrimination of uptake and killing, two related but disparate components of macrophage phagocytic activity.

A number of studies have suggested mechanisms by which PAF might potentially alter killing efficiency. Neutrophils, when primed with PAF, evidence enhanced superoxide synthesis and release in response to N-formylmethionyl-leucyl-phenylalanine or phorbol 12-myristate 13-acetate (PMA) (36). Moreover, pretreatment of rat KC with PAF has been found to induce nitric oxide synthase gene expression and protein synthesis, thereby increasing nitric oxide production in response to lipopolysaccharide stimulation (24). PAF also primes elastase release from PMA-stimulated neutrophils (36) and increases levels of antimicrobial polypeptides within human neutrophils (26, 27). All of these findings suggest potential mechanisms by which PAF, released from EC, might prime macrophages for subsequent yeast challenge and thereby enhance ICCA in our model.

TNF-alpha was another candidate mediator of the stimulatory effect, because it has been reported to upregulate phagocytosis in human monocytes (13, 37) and increase phagocytic killing of the Mycobacterium avium complex in human monocyte-derived macrophages (1). However, not only did our coculture system fail to generate measurable levels of TNF-alpha but also the inhibition of TNF-alpha activity with anti-TNF-alpha had no effect on the EC-mediated stimulation of phagocytic killing.

In summary, microvascular EC potentiate murine peritoneal macrophage phagocytic killing via the release of a soluble mediator. This physiologically relevant effect can be explained by the well-known generation and release of PAF by EC.


    FOOTNOTES

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

Address for reprint requests and other correspondence: G. B. Bulkley, Blalock 685, Johns Hopkins Hospital, 600 N. Wolfe St., Baltimore, MD 21287-4685 (E-mail: gbulkley{at}jhmi.edu).

Received 3 February 1999; accepted in final form 27 July 1999.


    REFERENCES
TOP
ABSTRACT
INTRODUCTION
MATERIALS AND METHODS
RESULTS
DISCUSSION
REFERENCES

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Am J Physiol Heart Circ Physiol 278(1):H269-H276
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