1 Bonfils Blood Center and Departments of 2 Pediatrics and 3 Surgery, University of Colorado School of Medicine, Denver, Colorado 80230; and 4 Department of Pediatrics, Section of Leukocyte Biology, Baylor College of Medicine, Houston, Texas 77030
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ABSTRACT |
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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 hypothesized that priming of PMNs
alters their reactivity such that a second priming agent causes PMN
activation and endothelial cell damage. PMNs were primed or not with
LPS and then treated with lyso-PCs, and oxidase activation and elastase
release were measured. For coculture experiments, activation of human
pulmonary microvascular endothelial cells (HMVECs) was assessed by
ICAM-1 expression and chemokine release. HMVECs were stimulated or
not with LPS, PMNs were added, cells were 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 on 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.
neutrophils; endotoxin; lysophosphatidylcholines
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INTRODUCTION |
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NEUTROPHILS (PMNs) are the most abundant phagocyte in circulation and are a vital part of host defense, especially against bacterial and fungal infections (11, 82). The normal function of PMNs involves a stepwise 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 nonoxidative methods. Importantly, the microbicidal functions of PMNs mostly occur in the tissues, and 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 endothelial cells (EC) and the increased surface expression of EC adhesion molecules that 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). Whereas 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 on
the PMN NADPH oxidase such that agents that augment the oxidative burst
to a subsequent stimulus 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 by-products 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 physiological 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). Because 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-pretreated 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 hypothesized 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, mimicking transfusion of a septic patient with stored blood, to determine whether LPS-primed PMNs could be activated by lyso-PCs, a second priming agent. In the second portion of this study, 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, which were allowed to settle and were then activated with lyso-PCs or vehicle, and the number of viable HMVECs was counted. The roles of PMN adhesion to vascular endothelium, chemokine release, oxidase activation, and degranulation were investigated in this coculture model of PMN-mediated HMVEC damage.
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MATERIALS AND METHODS |
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Unless otherwise specified, all reagents were purchased from
Sigma Chemical (St. Louis, MO). A Thermomax plate reader was purchased
from Molecular Dynamics (Menlo Park, CA). Plastic microplates, manufactured by Nunc, were obtained from Life Sciences Products (Denver, CO). HMVECs of pulmonary origin and all media and tissue culture reagents were purchased from the Clonetics division of BioWhittaker (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 intercellular adhesion molecule-1 (ICAM-1) were purchased from PharMingen (Torrey Pines, CA), and a fluorescein isothiocyanate (FITC)-labeled monoclonal antibody to CD54 was procured from Beckman Coulter (Miami, FL). A
monoclonal antibody to CD18 was obtained from Ancell (Bayport, MN).
Resveratrol and diphenyleneiodonium chloride (DPI) were obtained from Calbiochem-Novabiochem (San Diego, CA).
1,2-Bis(2-aminophenoxy)ethane-N,N,N',N'-tetraacetic acid-acetoxymethyl ester (BAPTA-AM) was purchased from Molecular Probes (Eugene, OR). Monoclonal antibodies to and ELISA kits for measuring epithelium-derived neutrophil-activating-78 (ENA-78), growth-related oncogene (GRO
), and IL-8 were obtained from R&D
Systems (Minneapolis, MN).
Lyso-PC 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-platelet-activating factor (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-min pulses by using a bath sonicator (model W-220F; Heat Systems-Ultrasonics, Plainview, NY) set at 30% maximal voltage. Lyso-PCs were tested at concentrations from 0.01 to 25 µM. Previous results demonstrated that higher concentrations of albumin (2-5%) actually further augmented the lyso-PC priming, nonspecific 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 pretreated for 30 min at 37°C with buffer control or LPS in
concentrations varying from 2 ng/ml to 2 µg/ml. Assays of oxidase
activation in response to lyso-PC or
N-formylmethionyl-leucyl-phenylalanine (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 previously (71, 74). The priming
activity of LPS was measured by first incubating the PMNs in the
reaction mixture containing LPS or Krebs-Ringer phosphate with 2%
dextrose (pH 7.35) (KRPD) control buffer for 3 min at 37°C,
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
O
Determination of elastase release in isolated PMNs. PMNs (1.5 × 106) were warmed to 37°C in a shaking water bath and then primed with 0.02-2 µg/ml LPS or buffer control for 5 min. The PMNs were activated with buffer, 0.45-14.5 µM lyso-PCs, or 1 µM fMLP as the positive control. After a 5-min reaction time, the PMNs were pelleted and the supernatant was 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 that the reduction of AAPVNA was secondary to that of elastase, we ran identical wells containing 5 µM of the specific elastase inhibitor methoxy-succinyl-alanyl-alanyl-prolyl-valyl chloromethyl ketone (AAPVCK) in conjunction with each treatment. Elastase release is reported as the percentage of total cellular elastase as determined by 0.1% Triton X-100 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/ml-2 µg/ml) for 2-12 h at 37°C,
7.5% CO2. The supernatants were aspirated, aliquoted, and
stored at
70°C for measurement of chemokine release. The adherent
HMVECs were removed with trypsin, washed, and incubated with an
FITC-labeled monoclonal antibody to ICAM-1 (CD54) for 30 min at 4°C
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 by employing enzyme-linked immunosorbent assay (ELISA) kits
purchased from R&D Systems.
IL-8 priming of NADPH oxidase.
PMNs were stimulated with IL-8
(1012-10
6 M) for 5 min at 37°C, and
superoxide anion production was measured as the maximal rate of
SOD-inhibitable reduction of cytochrome c at 550 nm of light as described previously (71, 74). PMNs were also incubated for 5 min at 37°C with IL-8
(10
12-10
9 M) and then activated with 1 mM fMLP, and the maximal rate of superoxide anion production was
measured as described above. The data (nmol
O
1 · min
1)
are expressed as means ± SE or as the relative 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/ml-2 µg/ml) and the other half with buffer for 6 h at 37°C, 7.5% CO2.
PMNs (1 × 106) were added, at a 10:1 effector
cell-to-target cell ratio, and allowed to settle for 30 min. After
settling, the PMNs were exposed to buffer, 200 ng/ml phorbol
12-myristate 13-acetate (PMA), or 0.45-14.5 µM lyso-PCs for 60 min. 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 EC 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 2 µg/ml LPS. Half of the wells received
PMNs incubated with 1 µg/ml CD18 for 10 min before their addition to
the HMVECs, whereas the other half received PMNs preincubated with
an isotypic control antibody. Notably, 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-1-mediated firm adhesion, we stimulated HMVECs with LPS for
6 h, and then 50% of the wells received 1 µg/ml of monoclonal antibodies to GRO
, ENA-78, IL-8, or ICAM-1 (CD54) for 10 min before
PMNs were added. The antibodies to the chemokines all had the ability
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 surface area
was counted to assess PMN-mediated EC cytotoxicity. In selected
experiments, an inhibitor of elastase, 5 µM AAPVCK, or intracellular
inhibitors of the oxidase, 1 µM resveratrol or 1-10 µM DPI,
were added either to the wells 30 s before the addition of PMNs or
to the PMNs 30 min before their addition to the coculture, 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 µM concentration of resveratrol and 1-10 µM concentrations of
DPI inhibited activation of the oxidase by 50-75% without
affecting cellular integrity. Finally, to block lyso-PC-mediated changes in cytosolic Ca2+ concentration, we loaded PMNs for
30 min with 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 h at 37°C, 7.5%
CO2. In selected wells, 1 mg/ml of neutralizing antibodies
or isotype controls to ENA-78, GRO
, and IL-8 were added 10 min
before the inclusion of PMNs. PMNs (1 × 106) were
then added and allowed to adhere for 60 min. 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 200 g for 5 min, and
the supernatant was discarded. The adherent cells were lysed with
0.01% Triton X-100, and the total amount of PMN elastase per well was
determined as mentioned previously and compared with the total cellular
elastase from the identical number of PMNs added to each well. The data
are the percentage of adherent PMNs expressed as means ± SE.
Statistical analysis. The means, SD, and SE 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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RESULTS |
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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 on these data, lyso-PC concentrations that primed the NADPH oxidase were employed
(0.45-14.5 µM). No concentrations of the lyso-PC mixture caused
activation of the PMN oxidase (albumin: 0.2 ± 0.2 vs. 14.5 µM;
lyso-PCs: 0.2 ± 0.2 nmol O
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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/ml to 2 µg/ml and lyso-PC concentrations from 0.45 to 14.5 µM (Fig. 2). Compared with
vehicle-treated controls, 0.45 µM lyso-PC did not cause any elastase
release, and this concentration was not augmented by pretreatment with
any concentration of LPS. In addition, concentrations of 2 ng/ml LPS
(Fig. 2) to 200 ng/ml LPS (results not shown) did not result in the
augmentation of elastase release in response to either lyso-PCs or
fMLP. At the 2 µg/ml concentration, LPS priming resulted in a direct
increase in the release of elastase by treatment of PMNs with 14.5 µM
lyso-PCs, but not 4.5 µM lyso-PCs, compared with both buffer-primed
controls and LPS-treated PMNs with buffer (Fig. 2). Lyso-PCs at 4.5 µM and 14.5 µM did cause increased amounts of elastase release
compared with albumin-treated controls (Fig. 2), similar to reports of other priming agents including PAF (80). Moreover, the
positive control, 1 µM fMLP, showed similar results.
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Activation of pulmonary HMVECs.
HMVECs were incubated with 2 µg/ml LPS from 2 to 12 h, and
the surface expression of ICAM-1 was measured by employing a FITC monoclonal antibody to ICAM-1 (CD54) and flow cytometry. Increases in
ICAM-1 began at 2 h (2.4 ± 0.6-fold increase,
P < 0.05 compared with media-treated controls) and at
6 h appeared similar to TNF--stimulated positive controls (LPS:
5.1 ± 0.6-fold increase vs. TNF-
: 6.7 ± 1.1-fold
increase, both compared with media-treated controls). At 12 h
there was a slight, but not statistically different, increase compared
with the 6-h incubation with LPS (6.7 ± 1.1 vs. 7.4 ± 1.5-fold increase compared with media-treated controls). Thus 6-h
incubations were used for all experiments to assess LPS activation of
HMVECs. The changes in ICAM-1 surface expression after 6 h of
LPS incubation (2 ng/ml-2 µg/ml) are shown in Fig.
3. LPS caused an increase in ICAM-1
surface expression at concentrations from 20 ng/ml to 2 µg/ml
(P < 0.05) but did not cause an increase in CD54 at an
LPS concentration of 2 ng/ml (Fig. 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 before the HMVECs were removed
(trypsin) from the 12-well plates. LPS concentrations from 20 ng/ml to
20 µg/ml were assessed for their ability to cause production and
release of ENA-78, 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 (Fig.
4). 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 control HMVECs at concentrations of
2-20 µg/ml LPS, although the amount of ENA-78 released at the 20 µg/ml concentration was not different from that at 2 µg/ml (Fig.
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
(Fig. 4B). Conversely, IL-8 was released maximally from the
LPS activation of HMVECs at an LPS concentration of 20 ng/ml;
higher concentrations of LPS did not augment the amount of IL-8
released by HMVECs (Fig. 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 µg/ml and higher caused release of significant
amounts of ENA-78 compared with media-stimulated controls.
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PMN-mediated HMVEC damage.
Pulmonary HMVECs were activated with buffer or LPS (2 ng/ml-2 µg/ml) for 6 h. 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 µM) and incubated for 60 min. 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 µg/ml) (Fig.
5,). Quiescent, buffer-treated HMVECs, incubated with lyso-PCs at 0.45-14.5
µM, did not display any evidence of killing of HMVECs;
similarly, quiescent, buffer-treated HMVECs incubated with PMNs
and lyso-PCs (0.45-14.5 µM) also exhibited no evidence of
killing. Furthermore, even the addition of 200 ng/ml PMA, a robust
activator of the NADPH oxidase, to PMNs coincubated with quiescent
HMVECs did not result in a decreased number of viable HMVECs
per 4 mm2 [HMVECs + PMNs: 1,166 ± 77 (mean + SE) vs. HMVECs + PMNs + PMA: 984 ± 159 HMVECs/4 mm2 (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). Also, importantly, 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 µg/ml when followed by the addition of PMNs
activated with 4.5 or 14.5 µM lyso-PCs (Fig. 5, A
and B). At an LPS concentration of 2 µg/ml, all
lyso-PC concentrations caused significant PMN-mediated HMVEC damage
compared with unstimulated HMVECs, LPS-activated HMVECs
coincubated with quiescent PMNs, and buffer-primed HMVECs treated
with PMNs and lyso-PCs (P < 0.05) (Fig.
5C). In addition, LPS-activated HMVECs incubated with
PMNs and treated with PMA also evidenced significant PMN-mediated
HMVEC cytotoxicity [HMVECs + PMNs: 1,166 ± 77;
HMVECs + PMNs + PMA: 984 ± 159; and HMVECs + 20 ng/ml LPS + PMNs + PMA: 520 ± 123 viable HMVECs/4 mm2 (P <0.05 compared with 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 (Fig. 5).
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Inhibition of PMN-mediated HMVEC damage with antibodies to
chemokines.
To determine the role of the chemokines released from LPS-activated
HMVECs, we added monoclonal antibodies to ENA-78, GRO, and IL-8
to the HMVEC reaction media after 6 h of LPS (2 µg/ml) stimulation before the addition of PMNs. As shown in Table
2, incubation with neutralizing,
monoclonal antibody to one or two chemokines attenuated the
PMN-mediated cytotoxicity compared with isotypic monoclonal antibody-
or media-treated controls. When antibodies to all three chemokines were
used, total abrogation of PMN-mediated HMVEC damage was observed.
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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 min before their addition to the coculture
to chelate the cytosolic Ca2+ and make it biologically
unavailable. As shown in Fig. 7, BAPTA chelation of cytosolic Ca2+ totally inhibited the
lyso-PC-mediated cytotoxicity of LPS-activated HMVECs, compared
with dimethyl sulfoxide (DMSO)-treated PMNs, which caused significant
HMVEC damage. Notably, upon visual inspection, BAPTA pretreatment
did not noticeably decrease the number of adherent PMNs compared with
PMNs preincubated with DMSO, although it abrogated HMVEC damage.
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DISCUSSION |
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TRALI 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 causes activation of the pulmonary endothelium, resulting in sequestration but not activation of host PMNs (70, 73, 74). Infusion of biological 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 cell damage, capillary leak, and pulmonary damage (70, 73, 74). Patients with acute infection or recent surgery may be predisposed to TRALI, and 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, intraperitoneal LPS, caused pulmonary leukostasis of PMNs, and the second insult, plasma and lipids from stored packed red blood cells 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 causes activation of the pulmonary endothelium,
causing increased surface expression of adhesion molecules and
chemokine release that prime PMNs, changing their phenotype to
adhesive, resulting in pulmonary vascular leukostasis, a
prerequisite for ALI (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, 9, 36, 38,
41, 42, 46, 51). However, the lung contains many small, tortuous
capillaries that may entrap rigid, primed PMNs, and such nondistensible
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 cell, capillary leak, and pulmonary injury
(11, 59, 64, 70, 74, 80). Acute lung injury, whether it be
TRALI or ARDS, is based on this two-event model (59, 62, 64, 74,
75, 84).
The data presented in this report confirm 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 on 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, because all three of
these agents are effective primers of the 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
pulmonary HMVECs. These studies demonstrated 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 Fig. 3 and as
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, only for PMNs that remain adherent when the plates are inverted
and subjected to 200 g for 5 min. 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
the addition of 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, preincubation of the PMN/HMVEC coculture with a selective elastase inhibitor, AAPVCK, did not affect PMN-mediated damage, data that are 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 suggest that oxidase activation is 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 without 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 physiological 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, 44, 45, 52, 57). Similar to PAF, lyso-PCs require
an albumin carrier and do not prime the PMN oxidase at concentrations
<0.45 µM (results not shown) (85). Moreover, the
addition of these compounds to fresh human plasma resulted in 1.7 ± 0.2-fold priming of the PMN oxidase compared with fresh plasma-pretreated controls (16, 25, 28, 29, 32, 37, 38, 40, 41, 47, 51, 54, 65, 68, 71 76, 84, 87).
In conclusion, one of the largest studies of ALI (70, 74) 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).
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ACKNOWLEDGEMENTS |
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This work was supported by a grant from The National Blood Foundation (to C. C. Silliman) and by National Heart, Lung, and Blood Institute Grants HL-59355-04 (to C. C. Silliman) and HL-42550 (to C. W. Smith). This paper was presented in part as a platform paper at The American Association of Blood Banks annual meeting, San Francisco, CA, Nov. 1999.
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FOOTNOTES |
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Address for reprint requests and other correspondence: C. C. Silliman, Bonfils Blood Center, 717 Yosemite Circle, Denver, CO 80230 (E-mail: christopher.silliman{at}uchsc.edu).
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. Section 1734 solely to indicate this fact.
July 24, 2002;10.1152/ajpcell.00540.2001
Received 13 November 2001; accepted in final form 19 July 2002.
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