A Two Insult In Vitro Model of PMN-Mediated Pulmonary Endothelial Damage: Requirements for CD18:ICAM-1 Adherence and Chemokine Release

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1A Two Insult In Vitro Model of PMN-Mediated Pulmonary Endothelial Damage:

Requirements for CD18:ICAM-1 Adherence and Chemokine Release

Travis H. Wyman1,2, A. Jason Bjornsen2, David J. Elzi1, C. Wayne Smith4, Kelly M. England2,

Marguerite Kelher2, Christopher C. Silliman1,2,3*

Bonfils Blood Center1 and the Departments of Pediatrics2 and Surgery3, University of Colorado

School of Medicine, Denver, CO and the Department of Leukocyte Biology4, Baylor College of

Medicine, Houston, TX.

*Correspondence: Christopher C. Silliman, MD, PhD

Associate Medical Director

Bonfils Blood Center

717 Yosemite Circle

Denver, CO 80230

Phone: (303) 363-2246

Fax: (303) 340-2616

E-mail: [email protected]

Running head: In vitro PMN-mediated endothelial damage

Copyright 2002 by the American Physiological Society.

AJP-Cell Articles in PresS. Published on July 24, 2002 as DOI 10.1152/ajpcell.00540.2001

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2 ABSTRACT

Lysophosphatidylcholines (lyso-PCs), generated during blood storage, are etiologic in a

two-insult, sepsis-based, model of transfusion related acute lung injury (TRALI). Individually

endotoxin (LPS) and lyso-PCs prime but do not activate neutrophils (PMNs). We hypothesize that

priming of PMNs alters their reactivity, such that a second priming agent causes PMN activation

and endothelial damage. PMNs were primed + LPS, treated with lyso-PCs and oxidase activation

and elastase release were measured. For co-culture experiments, activation of human pulmonary

microvascular endothelial cells (HMVECs) was assessed by ICAM-1 expression and chemokine

release. HMVECs were stimulated + LPS, PMNs were added, incubated with lyso-PCs, and the

number of viable HMVECs was counted. Lyso-PCs activated LPS-primed PMNs. HMVEC

activation resulted in increased ICAM-1 and release of ENA-78, GRO�, and IL-8. PMN-mediated

HMVEC damage was dependent upon LPS activation of HMVECs, chemokine release, PMN

adhesion, and lyso-PC activation of the oxidase. In conclusion, sequential exposure of PMNs to

priming agents activates the microbicidal arsenal, and PMN-mediated HMVEC damage was the

result of two insults: HMVEC activation and PMN oxidase assembly.

Key words: neutrophils, endotoxin, lysophosphatidylcholines, chemokines, endothelial damage.

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3INTRODUCTION

Neutrophils (PMNs) are the most abundant phagocyte in circulation and a vital part of host

defense, especially against bacterial and fungal infections (11;82). The normal function of PMNs

involves a step-wise progression of events that results in PMNs migrating from the circulation

through the vascular endothelium to the site of infection in the tissue (3;12;27;75). At the site of

infection, PMNs phagocytize the invasive microbes and kill them through both oxidative and non-

oxidative methods. It is important to note that the microbicidal functions of PMNs mostly occur in

the tissues, and that PMN priming by chemokines and other factors is part of the normal response to

infection (3;11;12;27;75;82). Priming of PMNs begins with their exposure to factors from

activated vascular endothelium, both chemokines released by activated EC and the increased

surface expression of EC adhesion molecules which initiate PMN adhesion, resulting in PMN

priming that may continue during chemotaxis to the inflammatory site (2;35;39;48). Primed PMNs

have enhanced microbicidal capacity to a subsequent stimulus so that microbial invaders may be

efficiently eradicated (2;35;39;48). While PMN priming is important for efficient killing of

bacteria and fungi, priming agents have been implicated in the pathogenesis of syndromes of PMN-

mediated organ damage including acute lung injury (ALI) (59;64;74;80).

Neutrophils are primed by a wide variety of stimulants that may be encountered during an

inflammatory response (1;2;15;16;30;49;71;80;88). Exposure to small concentrations of bacterial

endotoxin (lipopolysaccharide, LPS) is known to prime the respiratory burst and to augment, but

not cause elastase release from isolated PMNs (20-22;30). Priming is defined operationally upon

the PMN NADPH oxidase such that agents that augment the oxidative burst to a subsequent

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4stimulus, but do not individually cause oxidase assembly, are termed priming

agents(1;2;15;16;30;49;71;80;88). Priming agents are chemoattractants and affect other PMN

functions including changes in shape due to cytoskeletal rearrangements, firm adhesion mediated

by a conformational change in the �2-integrins, and the release of small amounts of granule

constituents (16;42;80). A number of compounds, including cytokines, the byproducts of the

complement cascade, and lipids, are priming agents and have been implicated in human disease;

however, many of the well described in vitro activators of the PMN oxidase, e.g. phorbol esters,

have little physiologic relevance or may never achieve concentrations in vivo that are employed

routinely in vitro (1;13;49;88). PMN priming agents have been shown to be etiologic in animal

models of ALI; however two priming agents must be administered sequentially (59;64;74).

Changes in PMN adherence, the enhanced release of cytotoxic products, and possible changes in

PMN reactivity due to the “primed” state have been proposed as contributing to tissue injury in

these conditions (70;80).

Previous studies have demonstrated that the routine storage of blood components, both

packed red blood cells and platelet concentrates, leads to the generation and accumulation of a

potent PMN priming activity, identified as a mixture of lysophosphatidylcholines (lyso-PCs)

(71;72). In addition, a number of investigators have shown that lyso-PC may augment the

respiratory burst in isolated human and rodent PMNs (13;19;26;71-74). Animal models of the

acute respiratory distress syndrome (ARDS) have postulated that two events are required;

moreover, animal models have employed the sequential administration of agents that have the

capacity to activate the vascular endothelium and prime the NADPH oxidase (11;59;64;74).

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5Because PMN priming agents have been implicated in ARDS, we postulated that the mixture of

lyso-PCs may act as a second insult and cause pulmonary damage in patients with transfusion

related acute lung injury (TRALI), a syndrome virtually identical to ARDS

(11;59;62;64;73;74;80). TRALI is thought to be secondary to the infusion of anti-leukocyte

antibodies that result in pulmonary sequestration of PMNs, activation of the complement

cascade, capillary leak and pulmonary injury, similar to ARDS (43;55;78;79). Because a number

of TRALI reactions did not have such an immune etiology, we postulated that TRALI, identical

to ARDS, is the result of at least two insults: the first is the clinical condition of the patient and

the second is the infusion of lyso-PCs in stored blood (9;10;73). A two-event animal model of

TRALI was developed, which demonstrated that the lungs from LPS pre-treated septic animals

developed acute lung injury in response to the plasma and lipids from stored but not fresh blood

products (74). Because of these findings we sought to determine the cellular physiology of

TRALI, and hypothesize that priming of PMNs alters their reactivity such that a normally

innocuous second agent activates the microbicidal arsenal of these primed PMNs culminating in

cytotoxicity. In the first portion of this study, isolated PMNs were primed with LPS and then

incubated with lyso-PCs, to mimic transfusion of a septic patient with stored blood, to determine

if LPS-primed PMNs could be activated by lyso-PCs, a second priming agent. To assess PMN-

mediated damage of human pulmonary microvascular endothelial cells (HMVECs), we

investigated LPS activation of these cells including increased surface expression of adhesion

molecules and chemokine release. Resting human PMNs were then added to both control and

LPS-treated HMVECs, allowed to settle, activated with lyso-PCs or vehicle, and the number of

viable HMVECs was counted. The roles of PMN adhesion to vascular endothelium, chemokine

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6release, oxidase activation, and degranulation were investigated in this co-culture model of

PMN-mediated HMVEC damage.

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7MATERIALS AND METHODS

Unless otherwise specified all reagents were purchased from Sigma Chemical Corporation,

St. Louis, MO. A Thermomax plate reader was purchased from Molecular Dynamics, Menlo Park,

CA. Plastic microplates manufactured by Nunc Inc., were obtained from Life Sciences Products,

Denver CO. Human microvascular endothelial cells of pulmonary origin (HMVECs) and all media

and tissue culture reagents were purchased from the Clonetics Division of BioWhittaker, Inc.,

Walkersville, MD. T-25 tissue culture flasks, 12 well plates, sterile pipettes, and paraformaldehyde

were obtained from Fisher Scientific, Pittsburgh, PA. A phycoerythrin-labeled monoclonal

antibody to CD11b and an unlabeled monoclonal antibody to ICAM-1 were purchased from BD

Pharmingen, Torrey Pines, CA, and a fluoroscein isothiocyanate labeled monoclonal antibody to

CD54 was procured from Beckman Coulter, Miami, FL. A monoclonal antibody to CD18 was

obtained form Ancell, Bayport, MN. Resveratrol and diphenyleneiodonium chloride (DPI) were

obtained from Calbiochem-Novabiochem Corp., San Diego, CA. 1,2-bis(2-aminophenoxy)ethane-

N,N,N′,N′–tetraacetic acid tetrakis(acetoxymethyl ester) (BAPTA) was purchased from Molecular

Probes, Eugene, OR. Monoclonal antibodies to and ELISA kits for measuring Epithelial-derived

Neutrophil Activating-78 (ENA-78), Growth Related Oncogene � (GRO��� and IL-8 were obtained

from R&D Systems, Minneapolis, MN.

Lysophosphatidylcholine preparation. The Lyso-PC mixture contained individual lyso-PCs

in the following molar ratios: 1-o-palmitoyl: 24, 1-o-oleoyl: 10, 1-o-stearoyl: 10, 1-o-hexadecyl

(C16) Lyso-PAF: 0.65, and 1-o-octadecyl (C18) Lyso-PAF: 0.35 (71). This mixture was solubilized

in 1.25% essential fatty acid free, globulin free human albumin with three 3 minute pulses using a

bath sonicator (model W-220F, Heat Systems-Ultrasonics, Inc., Plainview, NY) set at 30%

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8maximal voltage. Lyso-PCs were tested at concentrations from 0.01-25 µM. Previous results

demonstrated that higher concentrations of albumin, 2-5% actually further augmented the lyso-PC

priming, non-specific activity that precluded these albumin concentrations for solubilizing the lyso-

PCs (results not shown).

Neutrophil isolation and oxidase priming. PMNs were isolated by standard techniques

including dextran sedimentation, ficoll-hypaque gradient centrifugation, and hypotonic lysis of

contaminating red blood cells (71). Isolated PMNs were pre-treated for 30 minutes at 370C with

buffer control, or LPS in concentrations varying from 2 ng-2 µg/ml. Assays of oxidase activation in

response to lyso-PC or fMLP control were determined by measurement of the SOD-inhibitable

reduction of cytochrome c at 550 nm of light in a Thermomax Microplate Reader as described

(71;74). The priming activity of LPS was measured by first incubating the PMNs in the reaction

mixture containing LPS or KRPD control buffer for 3 minutes at 370C followed by activation of the

oxidase with the addition of lyso-PC. FMLP was used as a positive control for these experiments to

assess the integrity of the NADPH oxidase. Therefore, priming activity was measured as the

augmentation of the maximal rate of O2- in response to fMLP.

Determination of elastase release in isolated PMNs. PMNs (1.5x106) were warmed to 370C

in a shaking water bath and then primed with 0.02-2 µg/ml of LPS or buffer control for 5 minutes.

The PMNs were activated with buffer, 0.45-14.5 µM lyso-PCs or 1 µM fMLP as the positive

control. After a 5-minute reaction time, the PMNs were pelleted and the supernatant removed.

Elastase release was determined spectrophotometrically on the supernatant by the reduction of the

specific substrate methoxy-succinyl-alanyl-alanyl-prolyl-valyl p-nitroanilide (AAPVNA) at 405 nm

in duplicate. To ensure the reduction of AAPVNA was secondary to elastase, identical wells

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9containing 5 µM of the specific elastase inhibitor methoxy-succinyl-alanyl-alanyl-prolyl-valyl

chloromethyl ketone (AAPVCK) were run in conjunction with each treatment. Elastase release is

reported as the percentage of total cellular elastase as determined by 0.1% Triton-X paired

treatment of an identical number of PMNs.

HMVEC Activation. HMVECs were grown to >90% confluence on 12-well plates and

incubated with LPS [2 ng-2 �g/ml] for 2-12 hours at 37oC 7.5% CO2. The supernatants were

aspirated, aliquotted and stored at –70oC for measurement of chemokine release. The adherent

HMVECs were removed with trypsin, washed, and incubated with a FITC-labeled monoclonal

antibody to intercellular adhesion molecule-1 (ICAM-1, CD54) for 30 min at 4oC in the dark.

ICAM-1 surface expression was measured by flow cytometry. The supernatants were used for

direct measurement of ENA-78, GRO�, and IL-8 employing enzyme linked immunosorbent assay

(ELISA) kits purchased from R&D Systems, Minneapolis, MN.

IL-8 priming of the NADPH oxidase. PMNs were stimulated with IL-8 [10-12-10-6 M] for 5

min at 37oC and superoxide anion production was measured as the maximal rate of SOD-

inhibitable reduction of cytochrome c at 550 nm of light as described (71;74). PMNs were also

incubated for 5 min at 37oC with IL-8 [10-12 to 10-9 M] for 5 min and then activated with 1 mM

fMLP and the maximal rate of superoxide anion production was measured as described above.

The data (nmol O2-/ml/min) are expressed as the mean + the standard error of the mean or the fold

increase over buffer treated controls activated with fMLP.

HMVEC damage assay. HMVECs were grown to > 90% confluence in 12 well plates.

Half of the wells were incubated with LPS [2 ng-2 µg/ml] and the other half with buffer for 6 hours

at 370 C, 7.5% CO2. PMNs (1x106) were added, a 10:1 effector cell: target cell ratio, and allowed

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10to settle for 30 minutes. After settling the PMNs were exposed to buffer, 200 ng/ml phorbol

myristate acetate (PMA), or lyso-PCs [0.45-14.5 µM] for 60 minutes. The supernatants were

forcefully decanted by quickly inverting the plates onto absorbent towels and warm KRPD buffer

was added. The number of viable HMVECs, trypan blue negative, was counted over a 4 mm2

surface area by four separate observers, to exclude observer bias. Controls consisted of HMVECs

alone without PMNs. In addition, control HMVECs were also incubated with all of the reagents

used in these experiments alone or in combination. No single reagent or combination of reagents

caused HMVEC damage.

Inhibition of PMN-mediated EC damage with antibodies to CD18, ICAM-1, GRO�, ENA-

78, and IL-8 and inhibitors of the oxidase and PMN elastase. Inhibition of endothelial damage

from the context of the PMN was performed by growing HMVECs to > 90% confluence in 12 well

plates and then incubating all wells with LPS [2 µg/ml]. Half of the wells received PMNs

incubated with CD18 [1 �g/ml] for ten minutes prior to their addition to the HMVECs while the

other half received PMNs pre-incubated with an isotypic control antibody. It is of note that

incubation of these PMNs with this antibody to CD18 did not affect the oxidative burst of these

PMNs (results not shown). To block the effects of chemokines or ICAM-1mediated firm adhesion,

HMVECs were stimulated with LPS for 6 hours and then 50% of the wells received 1�g/ml of

monoclonal antibodies to GRO�, ENA-78, IL-8, or ICAM-1, CD54, for 10 minutes prior to the

addition of PMNs. The antibodies to the chemokines all had the capability to neutralize the

respective chemokines (R&D Systems). Similar to the CD18 experiments, the control HMVEC

wells received isotypic antibodies. The number of HMVECs per 4 mm2 was counted to assess

PMN-mediated EC cytotoxicity. In selected experiments, an inhibitor of elastase, 5 �M AAPVCK,

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11or intracellular inhibitors of the oxidase, 1 �M resveratrol or 1-10 �M diphenyleneiodonium

chloride (DPI), were either added to the wells 30 seconds prior to the addition of PMNs or to the

PMNs 30 minutes prior to their addition to the co-culture, respectively (63;67). The employed

concentrations of resveratrol and DPI were determined by inhibition of PMA-mediated oxidase

activation, and these inhibitors of the oxidase also effectively blocked superoxide anion production

to fMLP, PAF-primed PMNs stimulated with fMLP, and LPS-primed PMNs stimulated with lyso-

PCs (results not shown). The 1 �M concentration of resveratrol and concentrations of DPI from 1-

10 �M inhibited activation of the oxidase by 50-75% without affecting cellular integrity. Lastly, to

block lyso-PC-mediated changes in cytosolic Ca2+ concentration, PMNs were loaded for 30

minutes with 1,2-bis(2-aminophenoxy)ethane-N,N,N′,N′–tetraacetic acid tetrakis(acetoxymethyl

ester (BAPTA), a cell permeable, rapid chelator of cytosolic Ca2+, which has been demonstrated to

inhibit priming of the PMN oxidase (18).

PMN adherence to HMVECs. HMVECs were grown to >90% confluence in 12 well plates

and stimulated with buffer or LPS for 6 hours 370 C, 7.5% CO2. In selected wells 1mg/ml of

neutralizing antibodies, or isotype controls, to ENA-78, GRO�, and IL-8 were added 10 minutes

prior to the inclusion of PMNs. PMNs (1x106) were then added and allowed to adhere for 60

minutes. An aliquot of the identical number of PMNs was set aside. At the completion of

incubation, an adhesive covering was placed over the 12-well plates, the plates were centrifuged

inverted at 200g for 5 minutes, and the supernatant was discarded. The adherent cells were lysed

with 0.01% Triton X and the total amount of PMN elastase per well was determined as mentioned

previously and compared to the total cellular elastase from the identical number of PMNs added to

each well. The data is expressed as the mean + the standard error of the mean of the percentage of

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12adherent PMNs.

Statistical analysis. The mean, standard deviation and standard errors of the mean (SEM)

were calculated using standard techniques. Statistical differences among groups were determined by

a paired analysis of variance followed by a Tukey post hoc analysis for multiple comparisons.

Statistical significance was determined at the p<0.05 level.

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13RESULTS

Priming and activation of the PMN oxidase. Previous work demonstrated that the mixture

of lyso-PCs generated during routine blood storage was capable of priming the respiratory burst of

PMNs (71-74). Based upon these data, lyso-PC concentrations that primed the NADPH oxidase

were employed [0.45-14.5 �M]. No concentrations of the lyso-PC mixture caused activation of the

PMN oxidase, albumin control 0.2+0.2 versus 14.5 �M lyso-PCs 0.2+0.2 nmol O2-/min. To ensure

that the described priming activity was due to lyso-PCs and not platelet activating factor or like

compounds with an acetyl group in the sn-2 position, the lyso-PC mixture was incubated for 30

minutes at 37oC with 100 units/ml of either bee venom or active bovine pancreatic secretory

phospholipase A2 (sPLA2). The activity of the lyso-PC mixture was not diminished by sPLA2

treatment (results not shown). Figure 1 demonstrates that LPS-primed PMNs may be activated by

the addition of a second priming agent, lyso-PCs. This lyso-PC activation of the respiratory burst

was dependent upon both the LPS priming concentration and the “activating” concentration of lyso-

PCs. Buffer treated PMNs did not demonstrate activation of the PMN NADPH oxidase when

treated with any concentration of lyso-PCs; however, the fMLP positive control did cause

reproducible stimulation of the respiratory burst. LPS at 2 ng/ml did not cause priming of the

fMLP-activated respiratory burst nor did incubation of these PMNs with lyso-PCs result in oxidase

activation (Figure 1.). When the priming concentration of LPS was increased to 20 ng/ml, the 14.5

�M lyso-PC dose activated the NADPH oxidase. Moreover, this concentration of LPS also caused

significant priming of the fMLP-activated respiratory burst as compared to buffer- and 2 ng/ml

LPS-primed PMNs similar to previous reports (20-22;30). At 200 ng/ml of LPS, of LPS lyso-PC

concentrations of 4.5 and 14.5 �M activated the NADPH oxidase; furthermore, at an LPS

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14concentration of 2 �g/ml all concentrations of lyso-PCs [0.45-14.5 �M] caused oxidase activation.

In addition, LPS priming of the fMLP-activated respiratory burst was consistently increased over

the lower doses and became maximal at the 2 �g/ml LPS dose. It is of note that concentrations of

LPS < 2 ng/ml and lyso-PC < 0.45 µM displayed no evidence of priming or activation of the PMN

oxidase, respectively, and at all concentrations of LPS employed [2 ng-2 �g/ml], there was no

evidence of PMN lysis or loss of viability (PMNs were 99% viable by trypan blue exclusion).

Lastly, pre-treatment of PMNs with 50 �M BAPTA, an effective chelator of cytosolic Ca2+,

inhibited lyso-PC activation of the respiratory burst in LPS primed PMNs (92+5% - 97+3%) for all

concentrations of LPS and lyso-PCs employed.

Elastase release by PMNs. The ability of LPS to augment the lyso-PC elicited release of

elastase from isolated PMNs was evaluated over a range of LPS concentrations from 2 ng-2 µg/ml

and lyso-PC concentrations from 0.45-14.5 µM (Figure 2). As compared to vehicle-treated

controls, 0.45 �M lyso-PC did not cause any elastase release nor was this concentration augmented

by pre-treatment with any concentration of LPS. In addition, concentrations of 2-200 ng/ml of LPS

did not result in the augmentation of elastase release in response to either lyso-PCs or fMLP (Figure

2, results not shown). At the 2 �g/ml concentration, LPS priming resulted in a direct increase in the

release of elastase by treatment of PMNs with 14.5 �M lyso-PCs but not the 4.5 �M lyso-PCs as

compared to both buffer “primed” controls and LPS treated PMNs with buffer (Figure 2). Lyso-

PCs at 4.5 �M and 14.5 �M did cause increased amounts of elastase release as compared to

albumin-treated controls (Fig. 2) similar to reports of other priming agents including PAF (80).

Moreover, the positive control, 1 �M fMLP, showed similar results.

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15

Activation of human pulmonary microvascular endothelial cells. HMVECs were incubated

with 2 �g/ml LPS from 2-12 hours, and the surface expression of ICAM-1 was measured

employing a FITC monoclonal antibody to intercellular adhesion molecule-1 (ICAM-1, CD54) and

flow cytometry. Increases in ICAM-1 began at 2 hours (2.4+0.6-fold, p<0.05 as compared to

media-treated controls), and at six hours appeared similar to TNF�� stimulated positive controls

(LPS: 5.1+0.6-fold versus TNF�: 6.7+1.1-fold, both versus media treated control cells). At twelve

hours there was a slight, but not statistically different increase compared to the six-hour incubation

with LPS (6.7+1.1-fold versus 7.4+1.5-fold as compared to media-treated controls). Thus, six-hour

incubations were used for all experiments to assess LPS activation of HMVECs. The changes in

ICAM-1 surface expression following 6 hours of LPS incubation [2 ng-2 �g/ml] is illustrated in

Figure 3. LPS caused an increase in ICAM-1 surface expression at concentrations from 20 ng/ml to

2 �g/ml (p<0.05) but did not cause an increase in CD54 at an LPS concentration of 2 ng/ml (Figure

3). In addition, activation of endothelium, which is associated with chemokine release, was

measured in the incubation media (ELISA) from the same HMVECs employed for the ICAM-1

surface expression prior to removing (trypsin) the HMVECs from the 12-well plates. LPS

concentrations from 20 ng/ml to 20 �g/ml were assessed for their capability to cause production and

release of ENA78, GRO�, and IL-8. Concentrations of LPS that caused an increase in ICAM-1

surface expression also caused significant chemokine release for all three chemokines measured

(Figure 4, panels A-C). In these experiments the concentration response was taken an order of

magnitude higher to ensure that the HMVECs were “fully” stimulated by LPS for chemokine

production. The amount of ENA-78 released from HMVECs was statistically different from

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16control HMVECs at concentrations of 2-20 �g/ml of LPS, although the amount of ENA78 released

at the 20 �g/ml concentration was not different from the 2 �g/ml concentration (Figure 4A). The

release of GRO� became significant at LPS doses of 20 ng/ml and was maximal at 200 ng/ml;

higher concentrations of LPS did not induce an increased release of this chemokine from HMVECs

(Figure 4B). Conversely, IL-8 was released maximally from the LPS activation of HMVECs at an

LPS concentration of 20 ng/ml, for higher concentrations of LPS did not augment the amount of IL-

8 released by HMVECs (Figure 4C). Therefore, LPS concentrations that caused increased ICAM-1

surface expression also caused chemokine release of GRO� and IL-8, and only LPS concentrations

of 2 �g/ml and higher caused release of significant amounts of ENA78 as compared to media

stimulated controls.

To determine if the concentrations of chemokines caused priming or oxidase activation of

PMNs in vitro, PMNs were incubated with IL-8 for 5 min at 37oC and superoxide anion production

was measured. IL-8 [100 nM-1 �M] caused activation of the oxidase as compared to saline-treated

controls (nmol O2-/ml/min): saline, 0.1+0.5 IL-8 1 �M, 1.0+0.5*, 100 nM, 0.9+0.2*, 10 nM:

0.1+0.2, 1 �M fMLP 2.0+0.2*, n=6 (*=p<0.05 as compared to the saline-treated controls). At

concentrations of 1-10 nM IL-8 significantly primed the fMLP-activated respiratory burst by

2.5+1.0 to 6.2+2.0-fold (p<0.05 as compared to buffer primed fMLP-activated controls, n=8)and

lesser concentrations < 1 nM did not prime fMLP-activation of the PMN oxidase. Similar results

have been reported with respect to ENA-78 and GRO� (5;16;25;28;29;37;50;81). These data

provide supportive evidence that concentrations of the chemokines produced by LPS-activated

HMVECs, 1-5 ng/ml IL-8 < 1 nM, do not cause activation of the NADPH oxidase and are related

to priming of PMNs, which includes adherence.

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17

PMN-mediated HMVEC damage. Pulmonary HMVECs were activated with buffer or LPS

[2 ng-2 �g/ml] for 6 hours, PMNs were added, allowed to settle (30 min), and then activated with

buffer or lyso-PCs over a range of concentrations [0.45-14.5 �M] and incubated for 60 minutes.

The number of viable HMVECs, trypan blue negative, was counted over a 4 mm2 surface area. In

all cases, 99+2% of the adherent HMVECs were trypan blue negative; conversely, the detached

HMVECs in the supernatant were 99+4% trypan blue positive. LPS did not affect HMVEC

viability for any concentration employed [2 ng/ml-2 �g/ml] (Figure 5, panels A-C). Quiescent,

buffer-treated HMVECs, incubated with lyso-PCs 0.45-14.5��M did not display any evidence

killing of HMVECs, similarly, quiescent, buffer-treated HMVECs incubated with PMNs and lyso-

PCs [0.45-14.5 �M] also exhibited no evidence of killing. Furthermore, even the addition of 200

ng/ml of PMA, a robust activator of the NADPH oxidase, to PMNs co-incubated with quiescent

HMVECs did not result in a decreased number of viable HMVECs/4 mm2 (HMVECs + PMNs:

1166+77 (mean + SEM) versus HMVECs + PMNs + PMA: 984+159 HMVECs/4mm2, n=6). The

lowest dose of LPS, 2 ng/ml, did not result in any observed PMN-mediated cytotoxicity when

combined with lyso-PC activated PMNs for all lyso-PC concentrations tested (results not shown).

It is also important to note that lyso-PCs or PMA alone did not affect either quiescent HMVECs or

LPS-activated HMVECs in the absence of PMNs. However, PMN-mediated HMVEC damage

became readily apparent at LPS concentrations of 20 ng/ml - 2 �g/ml followed by the addition of

PMNs activated with 4.5 �M or 14.5 �M lyso-PCs (Figure 5, panels A and B). At an LPS

concentration of 2 �g/ml, all lyso-PC concentrations caused significant PMN-mediated HMVEC

damage as compared to unstimulated HMVECs, LPS-activated HMVECs co-incubated with

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18quiescent PMNs, and buffer primed HMVECs treated with PMNs and lyso-PCs (p<0.05) (Figure 5,

panel C). In addition, LPS activated HMVECs incubated with PMNs and treated with PMA also

evidenced significant PMN-mediated HMVEC cytotoxicity, HMVECs + PMNs: 1166+77,

HMVECs + PMNs + PMA: 984+159, HMVECs + 20 ng/ml LPS + PMNs + PMA: 520+123*

viable HMVECs/4mm2 (p<0.05 as compared to the other two groups, n=6). Thus, lyso-PC

activation of PMNs adhered to LPS-stimulated HMVECs resulted in destruction of HMVECs in a

concentration-dependent fashion (Figure 5).

To determine if firm adherence is required in this in vitro EC damage assay, PMNs were

pre-incubated with a monoclonal antibody to CD18 at saturating concentrations [1 �g/ml]. Pre-

incubation inhibited PMN-mediated HMVEC damage in HMVECs stimulated with 2 �g/ml LPS

and incubated with PMNs activated with 14.5 �M lyso-PCs (Figure 6, panel A, p<0.05, n=4).

Moreover, to investigate the role of EC adhesion molecules in this model, HMVECs were pre-

incubated (10 min) with a monoclonal antibody [1 �g/ml] to ICAM-1 (CD54). Similar to blocking

the PMN CD18 adhesion molecule, blockade of ICAM-1 also abrogated PMN-mediated HMVEC

damage in this model (Figure 6, panel B, p<0.05, n=4). These results indicate that firm adhesion

through �2-integrins on the PMN and ICAM-1 on the EC is required for PMN-mediated

cytotoxicity in this in vitro model of EC damage.

To characterize the cytotoxic agent responsible for the observed PMN-mediated HMVEC

damage, a selective elastase inhibitor, methoxy-succinyl-alanyl-alanyl-prolyl-valyl chloromethyl

ketone (AAPVCK), and two intracellular inhibitors of the NADPH oxidase, resveratrol and

C-00540-2001.R2

19diphenyleneiodonium chloride (DPI) were employed (63;67). To inhibit elastase, AAPVCK was

added to the reaction mixture 30 seconds prior to the addition of PMNs. Previous work with this

chloromethyl ketone has demonstrated that it does not inhibit the PMN oxidase or other signaling

pathways in PMNs (7). AAPVCK inhibition of PMN elastase had no effect on PMN-mediated

damage of LPS-primed HMVECs co-incubated with lyso-PC activated PMNs (results not shown).

Conversely, resveratrol and DPI inhibited PMN-mediated destruction of activated HMVECs (Table

1). Such inhibition of PMN-elicited cytotoxicity by resveratrol and DPI is strongly suggestive that

the oxidase played an important in this model.

Inhibition of PMN-mediated HMVEC damage with antibodies to chemokines. To

determine the role of the chemokines released from LPS activated HMVECs monoclonal antibodies

to ENA78, GRO�, and IL-8 were added to the HMVEC reaction media following 6 hours of LPS

[2 �g/ml] stimulation prior to the addition of PMNs. As illustrated in Table 2, incubation with

neutralizing, monoclonal antibody to one or two chemokines attenuated the PMN-mediated

cytotoxicity as compared to isotypic mAb- or media-treated controls. When antibodies to all three

chemokines were used, total abrogation of PMN-mediated HMVEC damage was observed.

Because the addition of neutralizing antibodies to the co-culture inhibited PMN cytotoxicity

in this two-event in vitro model we examined the possibility that these antibodies inhibited

chemokine mediated adhesion of PMNs to the activated HMVECs (Table 3). HMVECs were

incubated + 2 �g/ml LPS, the neutralizing antibodies were added to selective wells 10 min prior to

the addition of PMNs and then the PMNs were added and allowed to adhere for 60 min. Activated

HMVECs caused significant PMN adhesion that was abrogated by pre-incubation with the

C-00540-2001.R2

20chemokine antibodies employing an assay similar to previous published data (6). It is also

important to note that inverted centrifugation of HMVECs at 200g did not cause significant

detachment of these cells.

BAPTA inhibition of PMN cytotoxicity. Because changes in cytosolic Ca2+ concentration

are required for lyso-PC signaling in PMNs (69), PMNs were incubated with BAPTA for 30

minutes prior to their addition to the co-culture to chelate the cytosolic Ca2+ and make it

biologically unavailable. As shown in Figure 7, BAPTA chelation of cytosolic Ca2+ totally

inhibited the lyso-PC mediated cytotoxicity of LPS-activated HMVECs, as compared to dimethyl

sulfoxide (DMSO)-treated PMNs, which caused significant HMVEC damage. It is important to

note that upon visual inspection, BAPTA pre-treatment did not noticeably decrease the number of

adherent PMNs as compared to PMNs pre-incubated with DMSO, although it abrogated HMVEC

damage.

C-00540-2001.R2

21DISCUSSION

Transfusion related acute lung injury is identical to ARDS and is postulated to be the result

of infusing anti-leukocyte antibodies with transfusion of blood components (43;55). These anti-

leukocyte antibodies are directed against recipient antigens and cause pulmonary sequestration,

activation of the complement cascade, capillary leak, and pulmonary injury (43;55;78;79). Because

a number of observed TRALI reactions did not have an immune etiology, a two-event model was

proposed identical to animal models of ARDS (9;10;73;74). This model hypothesized that

susceptible patients must have an underlying clinical condition that cause activation of the

pulmonary endothelium, resulting in sequestration but not activation of host PMNs (70;73;74).

Infusion of biologic response modifiers, including lipids or even immunoglobulins directed against

specific granulocyte antigens, that activate these primed sequestered PMNs could then result in

activation of the microbicidal arsenal focused at the points of adherence, pulmonary endothelial

damage, capillary leak and pulmonary damage (70;73;74). Patients with acute infection or recent

surgery may be predisposed to TRALI and that the infusion of stored, but not fresh, blood with high

concentrations of lyso-PCs could cause activation of sequestered PMNs and pulmonary damage

(73). This hypothesis was tested in an animal model of TRALI in which the first insult, intra-

peritoneal LPS, caused pulmonary leukostasis of PMNs, and the second insult, plasma and lipids

from stored PRBCs including purified lyso-PCs solubilized in albumin, caused acute lung injury

(74). Because of these findings, we investigated the cellular physiology of the observed two-event

lung injury.

Prior work both in vitro and in animal models has postulated that the first event in ARDS

C-00540-2001.R2

22causes activation of the pulmonary endothelium causing increased surface expression of adhesion

molecules and chemokine release which primes PMNs changing their phenotype to from adhesive

resulting in pulmonary vascular leukostasis, a prerequisite for ALI which (8;11;14;29;37;46;74;75).

In this light, both endothelial cell adhesion molecules and their PMN ligands, �2-integrins, have

been reported to be essential in many models of PMN-mediated acute lung injury(8;41;46;51)

(9,36,38,42). However, it is also important to remember that the lung contains many small,

tortuous capillaries that may entrap rigid, primed PMNs and such non-distensible leukocytes would

then be unable to traverse the pulmonary vasculature, resulting in pulmonary leukostasis with points

of direct contact between the primed PMNs and the endothelium (17;31). The second insult causes

activation of the microbicidal arsenal of these adherent PMNs, which focuses the release of

cytotoxic agents at the points of PMN:endothelial cell adhesion or contact culminating in

endothelial damage capillary leak, and pulmonary injury (11;59;64;70;74;80). Acute lung injury

whether it be TRALI or ARDS is based upon this a two-event model (59;62;64;74;75;84).

The data presented in this report has confirmed that PMN priming not only causes adhesion

of PMNs to integrin consensus (RGD) ligands of activated HMVECs but also alters the reactivity

of PMNs such that these primed PMNs could be activated by the addition of a second priming

agent, lyso-PCs, in vitro (56). Moreover, neither LPS nor lyso-PCs given as single agents were able

to cause oxidase assembly, and activation of the oxidase and augmentation of elastase release were

dependent upon the concentrations of LPS, the first insult, and lyso-PCs, the second insult. In the

second portion of the study, the initial priming stimulus is not LPS but rather the chemokines

released as a function of HMVEC activation, for all three of these agents are effective primers of

C-00540-2001.R2

23the PMN oxidase and directly cause PMN adhesion presumably through a conformational change

in the �2-integrins (5;15;16;25;28;37;50;81). Thus, these data provide supportive evidence that the

exposure of PMNs to two sequential priming agents may activate their microbicidal arsenal and

cause PMN-mediated cytotoxicity.

The second part of this report examined PMN-mediated damage of human pulmonary

microvascular endothelial cells. These studies demonstrate that two sequential events or insults

may lead to PMN-mediated HMVEC damage in vitro. The first event consisted of LPS activation

of HMVECs resulting in increased surface expression of ICAM-1 and the release of chemokines

ENA-78, GRO�, and IL-8, which are effective PMN priming agents at the concentrations released

from HMVECs as shown in Figure 3 and demonstrated for IL-8 (5;15;16;25;28;37;50;81). These

chemokines were required for PMN adherence and together with firm adherence, a known priming

event, most likely altered the reactivity of these PMNs to a subsequent insult

(5;15;16;25;28;37;50;81). The addition of lyso-PCs, the second event, caused activation of these

primed PMNs that resulted in the focused release of cytotoxic agents, at the points of firm

adherence, that damaged and/or destroyed the activated HMVECs. Inhibition of PMN:HMVEC

adhesion with monoclonal antibodies to ICAM-1 (CD54) or CD18 abrogated PMN-mediated

HMVEC damage. Interruption of lyso-PC priming of the oxidase with BAPTA abrogated PMN-

mediated HMVEC damage by inhibiting activation of the oxidase and attenuating adherence.

Furthermore, the inclusion of neutralizing, monoclonal antibodies to all three chemokines

abrogated HMVEC damage in this model including the adherence of PMNs to activated HMVECs.

These adherence assays do not invite comparison to other leukocyte adherence assays for only

C-00540-2001.R2

24PMNs that remain adherent when the plates were inverted and subjected to 200g for 5 minutes.

Thus, only firmly adherent PMNs remained attached to the HMVECs. Incomplete inhibition of

PMN-mediated damage was demonstrated when a single neutralizing chemokine antibody or any

combination of two chemokine antibodies was added. Moreover, the addition of primed PMNs to

quiescent HMVECs or quiescent PMNs to activated HMVECs had no effect on HMVEC integrity,

but the latter group did elicit PMN adherence to the LPS-activated HMVECs. Without PMNs,

none of the stimuli employed alone or in combination affected HMVEC integrity including the

LPS/lyso-PC combination.

In addition, we investigated the components of the microbicidal arsenal responsible for

PMN-mediated damage of activated HMVECs. Although both elastase release and oxidase

activation could cause HMVEC damage, pre-incubation of the PMN/HMVEC co-culture with a

selective elastase inhibitor, AAPVPCK, did not affect PMN-mediated damage, data opposed to

other adherence-based killing of PMN targets, including previous work from this laboratory (6;83).

Conversely, inhibitors of the respiratory burst, both DPI and resveratrol, inhibited PMN mediated

HMVEC damage without affecting the cellular integrity of PMNs or the qualitative adhesion of

PMNs to activated HMVECs. Taken together, these data suggested that oxidase activation was

important for PMN-mediated HMVEC damage. Future experiments exploring the individual roles

of the chemokines released from activated HMVECs may provide more insight into the cellular

physiology of PMN-mediated HMVEC damage.

In the presented model, PMN-mediated HMVEC damage occurred in a static environment

C-00540-2001.R2

25without blood flow. Furthermore, pulmonary HMVEC activation resulted in increased surface

expression of ICAM-1 but not vascular cell adhesion molecule-1 (VCAM-1). Conversely, we were

able to show an increase in VCAM-1 on the surface of activated, human umbilical vein endothelial

cells (HUVECs) (results not shown). Other in vitro models of endothelial cell damage have

implicated VCAM-1 or other �2-integrin ligands, but many of these models employed flow

chambers or HUVECs that may have little physiologic relevance to the human lung

(7;23;36;60;61;77). Although previous work from this and other laboratories has demonstrated that

lyso-PCs can prime PMNs, a number of investigators have asserted that lyso-PCs are inactive with

respect to both leukocytes and platelets (4;33;34;44;45;52;53;57;58). Similar to PAF, lyso-PCs

require an albumin carrier and do not prime the PMN oxidase at concentrations less than 0.45 �M

(results not shown, (85). Moreover, the addition of these compounds to fresh, human plasma

resulted in priming of the PMN oxidase by 1.7+0.2-fold as compared to fresh plasma pre-treated

controls (71). (16;25;28;29;32;37;38;40;41;47;51;54;65;68;76;84;87)

In conclusion, (70;74)one of the largest studies of ALI demonstrated that blood transfusion

was the most commonly associated event; however, this transfusion requirement was deemed a

marker of clinical injury and not a possible etiology (24). In traumatically injured patients,

transfusion is a robust, independent predictor of the postinjury multiple organ failure syndrome

(MOF), which includes ALI (66). More importantly, the infusion of older stored blood, which

contains significant amounts of lyso-PCs, into trauma patients was also associated with the

development of ALI/MOF indicating that TRALI may be more common than previously reported

(70;86).

C-00540-2001.R2

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42ACKNOWLEDGEMENTS

This work was supported by a grant from The National Blood Foundation and grant number

#HL59355-04 from the NHLBI, NIH.

Presented in part as a platform paper at The American Association of Blood Banks annual meeting,

San Francisco, CA, November 1999.

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43FIGURE LEGENDS

Figure 1. Dose dependent activation of the PMN oxidase by sequential exposure to two

distinct priming agents. The concentration dependent lysophosphatidylcholine (L-PC) [0.45-14.5

�M] activation of the PMN oxidase is shown in relation to varying concentrations [2 ng/ml-2

�g/ml] of the LPS priming agent, and the data are expressed as the mean + the standard error of

the mean. PMNs were incubated with LPS for 30 minutes at 37oC and then stimulated with L-

PC, 1 �M fMLP, or buffer and the maximal rate of superoxide anion production was measured

(nmol/min). As the concentration of the LPS increases the concentration of the L-PC required to

activate the PMN oxidase subsequently decreases (* denotes statistical significance as compared

to buffer activated controls (p<0.05)). It is important to note that at an LPS concentration of 2

ng/ml there was no statistically significant increase of the oxidase response. FMLP was

employed as a positive control, and LPS doses of 20 ng/ml significantly primed this response

(p<0.05). This figure comprises data from seven separate experiments.

Figure 2. Elastase released from LPS-primed PMNs in response to L-PC stimulation.

Isolated PMNs were treated with buffer or LPS [20 ng/ml-2 �g/ml] for 30 min at 370C and

activated with buffer, L-PC {0.45-14.5 �M] or 1 �M fMLP (positive control). Total elastase

release was measured as the percentage elastase released into the supernatant in comparison to

triton-X lysed PMNs, and the data are depicted as the mean + the standard error of the mean. It

is important to note that statistical significance (*) (p<0.05) was attained only at an LPS

concentration of 2 �g/ml and a lyso-PC concentration of 14.5 �M. This figure represents the

results of seven separate experiments.

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44Figure 3. LPS-mediated increased surface expression of ICAM-1 on pulmonary

HMVECs. The surface expression of ICAM-1 on pulmonary HMVECs as quantified by flow

cytometry is represented as a function of the dose of endotoxin. The data are illustrated as the

mean + the standard error of the mean. Pulmonary HMVECs were treated with endotoxin

[0.002-2 �g/ml] or vehicle control (C) for 6 hours at 37oC 7.5% CO2. The supernatants were

aspirated and the cells were removed with trypsin and incubated for 30 minutes at 4oC with a

FITC-labeled monoclonal antibody to ICAM-1. The figure represents a sample size of eight, and

the * denotes statistical significance as compared to the vehicle-treated control pulmonary

HMVECs (p<0.05).

Figure 4. LPS-mediated release of chemokines from pulmonary HMVECs. Chemokine

release (panel A: ENA 78, panel B: GRO�, and panel C: IL-8) is represented as a function of

LPS concentration. The chemokines were measured in the media from HMVECs incubated for

six hours with differing doses of LPS [0.02-2 �g/ml] or vehicle control (C) by specific ELISA

assays in duplicate and calculated from a standard curve. The data, mean + standard error of the

mean, represent a sample size of 16, and the asterisks represent statistical difference from the

media of control HMVECs incubated for six hours with the saline (p<0.05). The # denotes

statistical differences from lower doses and the vehicle control (p<0.05, panel B).

Figure 5. PMN-mediated damage of pulmonary HMVECs. The damage caused by L-PC

activated PMNs on HMVECs primed with LPS at differing concentrations: A) 20 ng/ml LPS, B)

200 ng/ml LPS, and C) 2 �g/ml LPS stimulation for 6 hours at 37oC, 7.5% CO2. PMNs were

added, allowed to settle and stimulated with buffer or L-PCs [0.45-14.5 �M] for 60 minutes. The

amount of viable, adherent HMVECs, as determined by the ability to exclude trypan blue, was

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45counted via a dissecting microscope over a surface are of 4 mm2. Four different observers were

employed to alleviate observer bias, and the results, expressed as the mean + the standard error of

the mean, were within 2.4+2% of one another for all measurements. These damage assays

demonstrated dose dependence of both the first event, LPS, and the second event, L-PCs, for

PMN-mediated HMVEC damage (* denotes statistically significant HMVEC damage as

compared to HMVECs alone and HMVECs+ PMNs). In selected experiments the supernatants

were aspirated and stained and the discernible HMVECs were 99+4% trypan blue positive. This

figure is a representation of seven separate experiments for each lyso-PC concentration.

Figure 6. The effect of CD18 and ICAM-1 blockade on PMN-mediated pulmonary

HMVEC damage. The number of pulmonary HMVECs is represented as a function of treatment

group with pre-treatment of PMNs with a monoclonal antibody to CD18 (panel A) or pre-

incubation of HMVEC with a monoclonal antibody to ICAM-1 (panel B). All HMVECs were

incubated with 2 �g/ml LPS for 6 hours at 37oC 7.5% CO2 and selected wells had a monoclonal

antibody to CD18 (solid bars, Panel A) or ICAM-1 (solid bars panel B) followed by isolated

PMNs. The PMNs were allowed to settle for 30 min and selected wells were incubated with 14.5

�M lyso-PCs. The data, mean + standard error of the mean, are from four separate experiments

for both panels A and B, and the * in both panels denotes statistical difference from all other

treatment groups.

Figure 7. The effect of PMN pre-treatment with BAPTA on PMN-mediated HMVEC

damage. The number of pulmonary HMVECs per 4 mm2 is represented as a function of

treatment group. The HMVECs were treated with 2 �g/ml LPS for 6 hours at 37oC, 7.5% CO2.

PMNs were pre-treated with 50 �M BAPTA or DMSO control and then added to selected wells

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46of activated HMVECs, allowed to settle (30 min), and then certain wells received 14.5 �M lyso-

PCs. Following 60 min of incubation, the viable adherent HMVECs were counted per 4mm2.

These data, mean + standard error of the mean, represent a sample size of four, and the * denotes

statistical significance as compared to all treatment groups.

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47Table 1. Inhibition of PMN-mediated killing of LPS-activated pulmonary endothelial cells by

resveratrol and DPI.

Buffer PMNs PMNs+Lyso-PCs

DMSO 1088+22 1073+66 668+26*

RESVERATROL 1027+38 1033+41 885+24*#

DPI [10 �M] 1042+20 1012+44 968+38#

DPI [5 �M] 907+30*#

DPI [1�M] 670+42*

The data is expressed as the mean + standard error of the mean of viable HMVECs/4mm2 for

four separate experiments. Only the highest concentration of diphenyleneiodonium chloride

(DPI) was used for both buffer treatment and PMNs and HMVECs alone. *p<0.05 as compared

to buffer and PMNs alone. #p<0.05 as compared to the PMNs+lyso-PCs.

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48Table 2. Inhibition of PMN- mediated killing by monoclonal antibodies to chemokines.

Monoclonal antibody pre-treatment

Viable HMVECs/4mm2 Endotoxin PMNs Lyso-PCs Anti-ENA78 Anti-GRO� Anti-IL-8

1356+182 X

1325+159 X X

1355+169 X X

847+139* X X X

1113+130† X X X X

1095+172† X X X X

959+156† X X X X

1146+161† X X X X X

1140+160† X X X X X

1205+219† X X X X X

1332+177# X X X X X X

The data are expressed as the mean + the standard error of the mean of the number of viable pulmonary HMVECs/4mm2. * denotes

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49statistical significance as compared to all groups; † denotes statistical significance as compared to LPS activated HMVECs incubated

with lyso-PC activated PMNs, and # denotes statistical significance as compared to LPS activated HMVECs incubated with lyso-PC

activated PMNs and all single and double monoclonal antibody pre-treated HMVECs. This table summarizes the data from eight

identical experiments.

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50Table 3. Neutralizing antibodies to chemokines block PMN adherence to activated HMVECs.

Endothelial treatment Adherent PMNs (%)

Buffer 1.3+0.9

LPS 13.3+3.9*

LPS + antibodies 2.0+1.0#

The data are expressed as the mean + the standard error of the mean of four separate experiments

employing pulmonary HMVECs and the PMNs from four separate healthy donors. The *

denotes statistical significance as compared to the buffer controls and the # as compared to the

LPS activated HMVECs

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51ABBREVIATIONS

DPI = Diphenyleneiodonium chloride

ENA-78 = Epithelial-derived neutrophil activated 78

GRO� = Growth related oncogene ��

HMVEC = Human microvascular pulmonary endothelial cells

HUVEC = Human umbilical vein endothelial cells

Lyso-PCs = lysophosphatidylcholines

L-PC = lysophosphatidylcholines

AAPVNA = Methoxy-succinyl-alanyl-alanyl-prolyl-valyl p-nitroanilide

AAPVCK = Methoxy-succinyl-alanyl-alanyl-prolyl-valyl chloromethyl ketone

PAF = Platelet activating factor

PMN = neutrophil

sPLA2 = Secretory phospholipase A2

BAPTA = 1,2-bis(2-aminophenoxy)ethane-N,N,N′,N′–tetraacetic acid

tetrakis(acetoxymethyl ester)

Fig. 1

Fig. 2

Fig. 3

Fig. 4

A. B.

C.

Fig. 5A. B.

C.

Fig. 6

A.

B.

Fig. 7