Introduction

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Polymorphonuclear leukocytes (PMNs) play an important role in the pathogenesis of endothelial injury, such as acute respiratory distress syndrome (1). The initiating event in the development of the injury is PMN sequestration in microvessels (3). Factors that have been proposed to promote PMN sequestration are the size and deformability of PMNs (1), as well as the adhesive qualities of PMNs and endothelial cells (4).

Inflammatory stimuli increase PMN transit time by decreasing their deformability and increasing their adhesiveness to endothelial cells. The decrease in deformability is mediated by a rapid assembly of filamentous F-actin from soluble G-actin at the cell periphery, which increases the rigidity of PMNs and increases PMN transit time through the lung (5). PMN transit is also slowed by increased adherence between PMN and endothelial cells. This interaction between adhesion molecules on PMNs and their ligands on endothelial cells contributes to prolonged PMN sequestration in the lung (4, 8).

Recent findings suggest that endothelin (ET), a potent vasoconstrictive 21-amino acid peptide (9) released from endothelial cells, affects interaction between PMNs and the endothelium. Plasma ET-1 levels increase during sepsis, cardiac surgery, and acute respiratory distress syndrome (10). This suggests that ET-1 is produced in response to acute inflammatory stimuli, is an integral component of the systemic inflammatory reaction, and plays a role in the inflammatory process. Infusion of ET-1 causes PMN sequestration in rat lungs (13) and rabbit hearts (14). It also increases vascular permeability in the airways, gastrointestinal tract, and kidney in rats (15). Conversely, ET-1 receptor antagonist prevents PMN sequestration and injury in rat after an ischemia-reperfusion injury (16) and inhibits PMN infiltration in mice (17). The mechanisms of these effects of ET-1 on PMN-endothelial interaction are still unclear.

The vascular smooth-muscle constricting effect of endothelial-derived ET-1 and the opposing relaxing effect of nitric oxide (NO) are well studied (18). However, recent evidence indicates that ET-1 and NO influence the inflammatory response by modulating PMN-endothelial cell interactions (13, 19). Our working hypothesis is that ET-1 increases PMN sequestration by altering PMN deformability and adhesive properties. The present study was designed to evaluate the effect of ET-1 and NO on the factors that determine PMN sequestration in the lung. In in vitro studies, we measured the effects of ET-1 and NO on F-actin assembly, deformability, and adhesion molecule expression of PMNs; and in in vivo studies, we determined the retention of ET-1-treated PMNs in the lung.

	Materials and Methods

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In Vitro Studies

Cell preparation. Blood samples were collected from healthy volunteers (n = 6) in acid citrate dextrose as an anticoagulant. Leukocyte-rich plasma (LRP) was prepared by sedimenting red blood cells (RBCs) using 4% dextran (average molecular wt 162,000; Sigma Chemical Co., St. Louis, MO) in PMN buffer (138 mM NaCl, 2.7 mM KCl, 8.1 mM Na₂HPO₄ · 7H₂O, 1.5 mM KH₂PO₄, and 5.5 mM glucose, pH 7.4). LRP was centrifuged and residual RBCs were lysed by brief hypotonic shock with sterile water. PMNs were then separated from the mononuclear cells by centrifugation on Histopaque (Sigma), with a density of 1.077 g/ml at 150 × g for 13 min. The PMN purity was > 95% with a viability of 98%, as assessed by trypan blue exclusion.

F-actin content assay. Purified PMNs were resuspended in Hanks' balanced salt solution (HBSS) (1.3 mM CaCl₂, 5.0 mM KCl, 0.3 mM KH₂PO₄, 0.5 mM MgCl₂ · 6H₂O, 0.4 mM MgSO₄ · 7H₂O, 138 mM NaCl, 4.0 mM NaHCO₃, and 0.3 mM Na₂HPO₄; GIBCO BRL, Gaithersburg, MD) at a concentration of 2.0 ± 0.1 × 10⁶/ ml. PMNs were stimulated with ET-1 (Sigma) at different concentrations (10¹⁰ to 10⁷ M) for 5, 15, 30, 60, 180, and 300 s at 37°C. The reaction was stopped by fixation using 3% paraformaldehyde (PFA) for 30 min. After washing with phosphate-buffered saline (PBS), PMNs were simultaneously permeabilized and stained for 30 min in the dark at 37°C with a fresh mixture of 2 mM L--lysophosphatidylcholine, palmitol (Sigma), and 1 U/ml of BODIPY FL phallacidin (Molecular Probes, Inc., Eugene, OR). Cells were washed twice with PBS, and F-actin content was measured using a flow cytometer (Epics XL; Coulter Electronics, Hialeah, FL) and expressed as the mean fluorescent intensity of 5,000 cells. The change in F-actin content was expressed as the percentage of change from the baseline value.

To evaluate the effect of both secondary production of NO in PMNs and exogenous NO on F-actin assembly in PMNs induced by ET-1, PMNs were preincubated with either the NO synthase inhibitor N-nitro-L-arginine methylester (L-NAME) (1 mM) or the NO donor sodium nitroprusside (SNP) (100 µM) for 10 min, and then stimulated by ET-1 (10¹⁰ or 10⁷ M) for 5 and 30 s. These samples were processed in the same manner as described earlier.

To evaluate the effect of ET-1 on PMNs in whole blood, whole blood was incubated with ET-1 (10¹⁰ or 10⁷ M) at 37°C. Stimulated PMNs were fixed and permeabilized, and RBCs were lysed using Intraprep Permeabilizing Reagent (Beckman Coulter, Inc., Palo Alto, CA). Further, to determine the effect of ET-1 receptor antagonist, whole-blood samples were preincubated with 10 µM of the nonselective ETA and ETB receptor antagonist TAK-044 (Takeda Chemical Industries Ltd., Osaka, Japan) for 10 min and then stimulated with ET-1 (10⁷ M). These samples were stained for 30 min in the dark at 37°C with 1 U/ml of BODIPY FL phallacidin and processed in the same manner as described earlier.

Deformability assay. PMN deformability was assessed by measuring the pressure needed to pass PMNs through a polycarbonate filter with a uniform pore diameter of 5 µm (Poretics, Livermore, CA), which is a modification of the in vitro filtration system described by Lennie and colleagues (22) and Lowe (23). LRP was suspended in 0.5% albumin containing HBSS at a concentration of 1.07 ± 0.02 × 10⁶/ml and was filtered at a constant flow rate of 3 ml/min for 300 s using an infusion pump. The filtration pressure was measured upstream from the filter continuously by a pressure transducer (Validyne Engineering, Northridge, CA) and recorded every second by a computerized recording system. Four groups were studied: (1) LRP alone; (2) LRP plus ET-1 (ET-1, 10⁷ M, was added before filtration); (3) LRP plus ET-1 (ET-1, 10¹⁰ M, was added before filtration); and (4) LRP plus SNP plus ET-1 (LRP was incubated with 100 µM of SNP for 10 min and then stimulated with 10⁷ M of ET-1 before filtration).

Adhesion molecule assay. Changes in the surface expression of CD11b were measured using flow cytometry. LRP was centrifuged and the pellet was resuspended in HBSS before stimulation with ET-1 (10¹⁰ to 10⁷ M) for 3 min at 37°C. The reaction was stopped by fixing cells with PFA at 0.3% (final concentration). Cells were incubated for 10 min with either 1 mg/ml of phycoerythrin-conjugated mouse monoclonal antihuman CD11b antibody (DAKO Laboratories, Denmark), or phycoerythrin-conjugated mouse immunoglobulin G2a (DAKO). The erythrocytes were lysed for 60 s with Immuno-lyse and leukocytes were fixed with Immuno-fix (commercial kit from Coulter Clone, Coulter Immuno, Hialeah, FL). PMNs were identified using the typical forward- and side-scatter pattern and the expression of CD11b was measured as mean fluorescent intensity of 5,000 cells (Model Profile Epics 2; Coulter Electronics). The change of CD11b was expressed as the percentage of change compared with the baseline.

To determine the effect of NO on CD11b upregulation induced by ET-1, samples were preincubated with SNP (100 µM) for 10 min before the stimulation with ET-1 (10¹⁰ and 10⁷ M). These samples were processed in the same manner as described earlier.

In Vivo Study

Animals. Female New Zealand white rabbits (n = 6; weight = 3.9 ± 0.1 kg) were used in this study, and all of the experimental procedures were approved by the Committee on Animal Care of the University of British Columbia. The rabbits were anesthetized with ketamine hydrochloride (35 to 50 mg/kg intramuscularly) and xylazine (5 mg/kg intramuscularly). Catheters were placed in the superior vena cava through the right external jugular vein and the aortic root via the left carotid artery. Heparin (100 U/kg) was administered after surgery.

PMN and RBC labeling. PMNs and RBCs were simultaneously labeled using the fluorochrome DiI (Molecular Probes), which is a member of the lipophilic carbocyanine dye family and has been used as general membrane stain. A total of 8 ml of acid citrate dextrose anticoagulated blood was drawn from the central ear artery. This blood was washed with 10 ml PMN buffer and centrifuged at 1,000 rpm for 8 min. The pellet was resuspended in 40 ml of PMN buffer containing 2 µM of Cell Tracker CM-DiI (Molecular Probes) and was incubated for 5 min at 37°C and then for 15 min at room temperature. Flow cytometry using typical forward- and side-scatter characteristics of cells were used to identify labeled cells. To eliminate possible overlap with monocytes, gates for PMN analysis were set on the larger and more granular cells in the PMN population. Preliminary experiments showed that this labeling procedure results in optimum labeling efficiency (93 ± 1% of PMNs and 14 ± 2% of RBCs). Further, this labeling procedure requires minimal cell manipulation and does not change PMN activation markers such as L-selectin or CD18 expression. Also, labeling human PMNs with DiI has been shown not to activate PMNs, which makes this labeling technique suitable to study PMN functions (24).

PMN transit through the lungs. Two indicator-dilution runs were carried out in the same animal after bolus injection of DiI-labeled RBCs and PMNs (2.5 ml of labeled blood was rapidly [1 s] injected into the superior vena cava). The first indicator dilution was done with naive labeled cells and was followed 30 min later with an injection of DiI-labeled cells that were incubated with ET-1 (10⁷ M) for 15 s ex vivo. Blood samples were collected from the aortic root catheter into preweighed tubes at 0.5-s intervals using a fraction collector for a 12-s period (point of recirculation) (25). The number and percentage of DiI-labeled PMNs in each sample were determined using flow cytometry after lysing the RBCs with Immuno-lyse and fixation with Immuno-fix (Coulter Clone, Coulter Electronics). The number of DiI-labeled PMNs in each sample was determined using flow cytometry and expressed as a fraction of the input value. The number of DiI-labeled RBCs was also determined using flow cytometry after diluting the sample in PBS 100 times. These numbers were expressed as fractions of input values. The cardiac output was determined using DiI-labeled RBCs as a flow indicator (26). The recovery of DiI-labeled PMNs into aortic root samples during their first passage (up to the recirculation of RBCs) was determined by integrating the curves and compared with the RBC curves. Retention of PMNs on their first passage through the lung was determined by subtracting recoveries from input values (25).

Statistics

F-actin content and adhesion molecules of PMNs were analyzed using a randomized block design analysis of variance (ANOVA), with donor as a blocking factor. The pressure data of the filtration study was analyzed using a randomized block design ANOVA for the areas under the curve, with donor as a blocking factor. The retention of DiI-labeled PMNs was analyzed using a one-way ANOVA. The sequential rejective Bonferroni test was used to correct for multiple comparisons (27). A corrected P value < 0.05 was considered significant. All values are expressed as means ± standard error.

	Results

Top Abstract Introduction Materials and Methods Results Discussion References

In Vitro Studies

F-actin content assay. ET-1 (10⁸ to 10⁷ M) caused a rapid increase in purified PMN F-actin content (with 10⁷ M of ET-1, at 5 s: 58 ± 6% increase from baseline, P < 0.01) that returned to baseline within 300 s (Figure 1) in a dose-dependent fashion. Lower concentrations of ET-1 (10¹⁰ to 10⁹ M) caused a decrease in purified PMN F-actin content (with 10¹⁰ M of ET-1, at 5 s: 6.9 ± 1.5% decrease from baseline, P < 0.01) that returned to baseline within 300 s (Figure 1).

View larger version (20K): [in this window] [in a new window]   Figure 1.   Effect of ET-1 on F-actin assembly of purified PMNs as measured by flow cytometry. PMNs were stimulated with ET-1 at different concentrations (1010, 109, 108, and 107 M). F-actin content immediately increased with stimulation from higher concentrations of ET-1 (108 and 107 M) and returned to baseline within 300 s. F-actin content decreased with stimulation from lower concentrations of ET-1 (1010 and 109 M) and returned to baseline within 60 s. Results are expressed as means ± standard error of the mean (SEM) of six experiments. *P < 0.01 versus baseline, **P < 0.05 versus baseline.

Preincubation of purified PMNs with L-NAME did not block the F-actin increase observed with high concentration of ET-1 (10⁷ M) (Figure 2A), but prevented the F-actin content decrease with low concentration of ET-1 (10¹⁰ M) (Figure 2B). Preincubation of purified PMNs with SNP inhibited the F-actin content increase observed with high concentration of ET-1 (10⁷ M) (P < 0.01; Figure 2A), and enhanced the F-actin content decrease observed with low concentration of ET-1 (10¹⁰ M; P < 0.01; Figure 2B).

View larger version (18K): [in this window] [in a new window]   Figure 2.   Effect of preincubation of purified PMNs with L-NAME or SNP on F-actin assembly of PMNs induced by ET-1 (A: 107 M, B: 1010 M). L-NAME did not change the F-actin content increase by 107 M of ET-1 (A), but prevented the F-actin content decrease by 1010 M of ET-1 (B). Preincubation of PMNs with SNP inhibited the F-actin content increase by 107 M of ET-1 and enhanced the F-actin content decrease by 1010 M of ET-1. Results are expressed as means ± SEM of six experiments. *P < 0.01 versus baseline, **P < 0.05 versus baseline.

The F-actin change of PMNs in whole blood by ET-1 (10¹⁰ and 10⁷ M) was not transient and continued for 300 s (Figure 3). The nonselective ETA and ETB receptor antagonist TAK-044 inhibited the F-actin increase of PMNs stimulated by ET-1 (10⁷ M; Figure 3).

View larger version (19K): [in this window] [in a new window]   Figure 3.   Effect of ET-1 on F-actin assembly of PMNs in whole blood with or without incubation with the nonselective ETA and ETB receptor antagonist TAK-044. ET-1 (1010 and 107 M) increased the PMN F-actin content for up to 300 s. TAK-044 inhibited the F-actin increase of PMN stimulated by ET-1 (107 M). Results are expressed as means ± SEM of six experiments. *P < 0.01 versus baseline, **P < 0.05 versus baseline.

Deformability assay. The pressure required to pass LRP through 5-µm-pore polycarbonate membrane filters increased when cells were preincubated with 10⁷ M of ET-1 (6.6 ± 0.4 to 8.9 ± 0.6 cm H₂O at 300 s; P < 0.01; Figure 4). Preincubation of LRP with SNP (100 µM) inhibited this increase in filtration pressure induced by 10⁷ M of ET-1. Incubating cells with low concentration of ET-1 (10¹⁰ M) or a combination of ET-1 (10¹⁰ M) and SNP (100 µM) did not change the filtration pressure.

View larger version (17K): [in this window] [in a new window]   Figure 4.   Effects of ET-1 and NO on the deformability of PMNs. LRP was filtered through 5-µm-pore polycarbonate membrane filters, and filtration pressure was measured over time. The pressure increased with 107 M of ET-1. Preincubation of LRP with SNP (100 µM) inhibited this increase. The amount of 1010 M of ET-1 did not change the filtration pressure. Results are expressed as means ± SEM of six experiments. *P < 0.01 LRP plus ET-1 versus LRP, **P < 0.05 LRP plus SNP plus ET-1 versus LRP plus ET-1 (107 M).

Adhesion molecule assay. ET-1 (10¹⁰ to 10⁷ M) increased the CD11b expression in a dose-dependent fashion (Figure 5A). The CD11b expression of PMN increased by 47 ± 15% after stimulation with 10⁷ M ET-1 (P < 0.01). Preincubation of PMN with SNP (100 µM) inhibited this increase of CD11b expression (P < 0.01; Figure 5B).

View larger version (18K): [in this window] [in a new window]   Figure 5.   (A) Effect of ET-1 on CD11b expression of PMNs as measured by flow cytometry. PMNs were stimulated with ET-1 at different concentrations (1010 to 107 M). The expression of CD11b was measured as mean fluorescent intensity of 5,000 cells. The change in CD11b was expressed as the percentage changes compared with the baseline. ET-1 increased the CD11b expression in a dose-dependent fashion. Results are expressed as means ± SEM of six experiments. *P < 0.01 versus baseline. (B) Effect of SNP on the increase in CD11b expression of PMNs induced by ET-1 (107 M to 1010 M). Preincubation of PMNs with SNP (100 µM) inhibited CD11b upregulation induced by ET-1. Results are expressed as means ± SEM of six experiments. *P < 0.01 versus baseline, **P < 0.01 versus ET-1 (107 M).

ET-1 (10¹⁰ to 10⁷ M) did not change L-selectin expression of PMN (data not shown).

In Vivo Study

First RBC transit through the lung. The fraction of DiI-labeled RBCs in aortic root samples compared with input value did not change with ET-1 treatment on their first transit through the lung (Figure 6). The cardiac output measured using DiI-labeled RBCs as a flow indicator did not change with ET-1 treatment (control group: 402 ± 25 ml/min, ET-1-treated group: 396 ± 25 ml/min). Blood loss was 5.6 ± 0.4 ml/run and replaced with normal saline with blood pressures unchanged during the experiment.

View larger version (17K): [in this window] [in a new window]   Figure 6.   The percentage of DiI-labeled RBCs in the aortic root samples. The fraction of DiI-labeled RBCs in aortic root samples did not change with ET-1 treatment. The cardiac output measured using DiI-labeled RBCs as a flow indicator did not change with ET-1 treatment (control group: 402 ± 25 ml/min; ET-1- treated group: 396 ± 25 ml/min). Results are expressed as means ± SEM of six experiments.

First PMN transit through the lung. The fraction of DiI-labeled RBCs and PMNs in the aortic root samples compared with input values of both groups are shown in Figure 7. The recovery of all the DiI-labeled PMNs from the aortic root samples was calculated and compared with the recovery of DiI-labeled RBCs (assumed as 100%). The retention of PMNs on their first passage through the lung increased from 45 ± 8 to 70 ± 5% with ET-1 treatment (P < 0.05).

View larger version (22K): [in this window] [in a new window]   Figure 7.   The percentage of DiI-labeled RBCs and PMNs in aortic root samples with (A) injection of naive cells or (B) injection of ET-1 (107 M)-treated cells. The recovery of DiI-labeled PMNs in the aortic root samples was calculated (see MATERIALS AND METHODS) and compared with the recovery of DiI-labeled RBCs. The retention of PMNs on their passage through the lung increased from 45 ± 8 to 70 ± 5% with ET-1 treatment (P < 0.05). Results are expressed as means ± SEM of six experiments.

	Discussion

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Sequestration of PMNs into microvessels is the first and an essential step in the recruitment of PMNs into a site of inflammation. In this study we determined the effect of ET-1 on the factors that regulate PMN sequestration in microvessels. In vitro studies showed that ET-1 ( 10⁸ M) caused a rapid and dose-dependent increase in F-actin content and a decrease in PMN deformability (Figure 4). The inhibition of these F-actin changes induced by ET-1 and blocked by the ETA and ETB blocker TAK-044 (Figure 3) clearly show this effect to be ET-1-specific. Changes in F-actin and deformability induced by ET-1 were inhibited by preincubation of PMNs with the NO donor SNP. These results suggest that ET-1 is an important mediator influencing PMN sequestration in lung microvessels because changes in PMN deformability have been shown to be the major factor contributing to PMN sequestration in the pulmonary vascular bed (2, 5).

Interestingly, lower concentration of ET-1 ( 10⁹ M) decreased the F-actin content of PMNs. This is similar to the dose-dependent effect of ET-1 on smooth-muscle contraction (28, 29) where higher concentration ( 10⁸ M) causes smooth-muscle constriction and lower concentration of ET-1 ( 10⁹ M) induces smooth-muscle relaxation. The relaxation by lower concentration of ET-1 is considered to be mediated by the secondary production of endogenous NO (18). The decrease in F-actin content induced by low concentration of ET-1 was inhibited by L-NAME and enhanced by SNP (Figure 2B), which indicates a similar mechanism with endogenous NO production in PMNs stimulated by the exposure to low concentrations of ET-1. The low endogenous NO production capability of PMNs is compatible with the inability of L-NAME to block the effect of high but not low concentrations of ET-1. In the filtration studies we could not show that low concentration of ET-1 changed leukocyte deformability significantly. We suspect that our method of measuring PMN deformability is not sensitive enough to detect the small changes in PMN deformability caused by the small decrease in F-actin induced by low concentration of ET-1. Further, we used LRP to perform the deformability assay to avoid PMN purification and activation. Changes in PMN deformability are the predominant factor responsible for pressure changes in our system using both rabbit (5) and human mixed leukocyte populations with different stimuli (30). Although we cannot exclude a differential effect of ET-1 on other leukocytes, such as monocytes, these cells most likely have a small influence on the results.

ET-1 upregulated CD11b expression on PMNs in a dose-dependent fashion (Figure 5A), which supports work by Lopez Farre and colleagues (14). We have extended these observations by demonstrating that this increase in PMN CD11b expression induced by ET-1 can be blocked by SNP (Figure 5B). ET-1 also upregulates the expression of intercellular adhesion molecule-1, vascular cell adhesion molecule (VCAM)-1, and E-selectin on endothelial cells (31), and NO reduces the expression of VCAM-1 on activated endothelium (32). Together, these findings suggest that ET-1 and NO are important mediators regulating adhesion molecules on PMNs and endothelium, thereby influencing PMN-endothelial adhesive interactions.

ET-1 induces a rapid increase in intracellular Ca²⁺ in human PMNs (33). This reaction plays an important role in signal transduction during PMN activation by activating cellular kinases and phosphatases, a key step in integrin upregulation and possibly in F-actin assembly (34, 35). Therefore, it is reasonable to speculate that changes in intracellular Ca²⁺ induced by ET-1 could be an important trigger inducing the CD11b upregulation and F-actin increase.

NO controls vascular tone by activating guanylate cyclase and increases the concentration of cyclic guanosine monophosphate (cGMP) in the smooth-muscle cell (36). We have previously shown that NO prevents the activation of PMNs stimulated by complement fragments (21). We suspect that NO also prevents the activation of PMNs caused by ET-1, because cGMP is an inhibitory secondary messenger in PMNs (37) that prevents the increase of intracellular Ca²⁺ by inhibiting Ca²⁺ influx through the Ca²⁺ channel (38). In addition, NO also promotes adenosine diphosphate ribosylation of actin that inhibits F-actin assembly (39). Our in vitro studies showing that NO inhibited PMN F-actin assembly, deformability, and CD11b expression induced by ET-1 could be via these mechanisms.

The plasma concentration of ET-1 is low in normal individuals (1 pg/ml, 4 × 10¹³ M) (18), and increases during endotoxemia (100 pg/ml, 4 × 10¹¹ M) (40). These circulating levels are insufficient to elicit vasoconstriction or cause the functional changes in PMNs demonstrated in these experiments. This suggests that ET-1 acts locally in a paracrine fashion rather than as a circulating mediator (18). The concentration of ET-1 at the interface between endothelial cells and intravascular PMNs is not known. We suspect that the local presence of ET-1 during the inflammatory response influences PMN endothelial interactions, and we speculate that ET-1 acts on cells such as PMNs that come in close contact with the endothelium during their transit through restricted vascular beds such as the pulmonary capillaries. This close contact provides the opportunity for ET released from endothelial cells to act directly on intravascular cells.

Several studies from our own (5) and other (2) laboratories have shown that a decrease in PMN deformability is the major factor responsible for rapid but reversible sequestration of PMNs in the lung. This is in contrast to adhesion molecules such as L-selectin and 2 integrins that cause a delayed but prolonged PMN sequestration (8, 41). The in vivo studies reported here show that PMNs treated with ET-1 are retained more in the lung compared with naive PMNs (Figure 7). The method used to label PMNs is novel in that it involves minimal cell manipulation during the labeling procedure. The fluorescent dye used to label the cells forms a stable bond with the cell membrane, and labeled cells are detected using flow cytometry. Although the cardiac output did not change when ET-1-treated blood was injected, the retention of PMNs on their first passage through the lung increased from 45 ± 7 to 70 ± 5%. This retention of naive PMNs is less than previously reported using radioisotopes to label PMNs (25). We suspect that PMN purification and labeling procedures used in these previous experiments mildly activated PMNs, resulting in more PMN retention. This is supported by our finding that the effects of ET-1 on PMN F-actin content changes in whole blood (Figure 3) were prolonged compared with the purified PMNs (Figure 1). We suspect that less manipulation of PMNs or the existence of plasma could produce a prolonged effect shown in the whole-blood study. Our in vivo results suggest that changes in PMN deformability induced by ET-1 resulted in the increase in PMN retention in the lung.

In summary, ET-1 changes the F-actin content of PMNs in a dose-dependent fashion. High concentrations of ET-1 increase F-actin content of PMN and decrease PMN deformability. These changes are inhibited by the NO donor SNP. In contrast, low concentrations of ET-1 cause a small but significant decrease in PMN F-actin content, most likely via endogenous NO production. ET-1 also increases the expression of CD11b on PMNs, which was inhibited by the NO donor SNP. Our in vivo studies showed that PMN treated with ET-1 were retained in the lung to a greater degree than naive PMNs. Together these studies suggest that ET-1 and NO are important in regulating the factors that determine PMN sequestration. We speculate that blocking ET receptors or exogenous NO could decrease PMN sequestration and PMN-mediated endothelial injury.