Pathogen-free male Wistar rats weighing 180 to 220 g were obtained from Nihon SLC (Hamamatsu, Japan). Cepharanthine was obtained from Kaken Syoyaku Co. (Tokyo, Japan). Bovine serum albumin, cytochalasin B, formyl-Met-Leu-Phe (fMLP), and nitrogen mustard (NM) were purchased from Sigma (St. Louis, MO), lipopolysaccharide (Escherichia coli, serotype 055:B5) from Difco (Detroit, MI), and Bolton-Hunter reagent and chromium 51 (⁵¹Cr) from Amersham International (Buckinghamshire, UK). All other reagents used were of analytic grade.

Animal model of LPS-induced acute pulmonary vascular injury. The study protocol was approved by the Kumamoto University Animal Care and Use Committee, and the care and handling of the animals were in accordance with the National Institutes of Health guidelines. Adult pathogen-free male Wistar rats (body mass, 180 to 220 g) were injected intravenously with 200,000 cpm/kg body weight of ¹²⁵I-labeled bovine serum albumin prepared with Bolton-Hunter reagent 5 min before intravenous administration of a bolus dose (5 mg/kg) of LPS into the tail vein. Cepharanthine (10 mg/kg) was administered intraperitoneally 30 min before injection of LPS. Negative control animals received saline instead of cepharanthine. Six hours after LPS administration, animals were anesthetized by intraperitoneal injections of pentobarbital sodium (50 mg/kg) and exsanguinated via the abdominal aorta. Blood samples were collected in tubes containing a 1/10 volume of 3.8% (wt/vol) sodium citrate. Blood was centrifuged at 2,000 × g for 10 min. The lung vasculature was perfused through the right cardiac ventricle with 10 ml of cold 0.9% NaCl. The lungs were then removed and the amount of radioactivity remaining in the tissue was measured with a gamma scintillation counter (Model 5130; Packard Instrument, Downers Grove, IL). LPS-induced pulmonary vascular damage was assessed in terms of the increase in vascular permeability, expressed as the permeability index, as previously described (11). The permeability index represents the ratio of the amount of radioactivity present in lung tissue to the amount of radioactivity present in 1 ml of blood obtained at the time of death. LPS-induced lung hemorrhage and pulmonary congestion were assessed by measuring the accumulation of ⁵¹Cr-labeled red blood cells in control and LPS-treated rats according to a previously described method (12). Animals received intravenous infusions of ⁵¹Cr-labeled red blood cells (50 μl containing 80,000 cpm) 30 min before injection of LPS. Rats were killed 6 h after administration of LPS. Lung radioactivity was measured in the presence and absence of saline perfusion and compared with the radioactivity present in blood. Bronchoalveolar lavage (BAL) was performed as described previously (11). In brief, after intraperitoneal administration of pentobarbital sodium, an incision was made in the anterior neck, and a catheter was secured in the trachea with a surgical suture. BAL was performed with 5 ml of saline. Radioactivity in the BAL fluid was measured with a gamma scintillation counter. There was no significant increase in the ratio of ¹²⁵I-labeled bovine serum albumin concentration in the BAL fluid to that in blood 6 h after administration of LPS (nearly zero), suggesting that the lung injury in this model was limited to the vascular endothelial cells (10).

Because the changes in permeability were small and there was no significant increase in the ratio of ¹²⁵I-labeled bovine serum albumin concentration in the BAL fluid to that in blood 6 h after administration of LPS, the water content of the lung was estimated. The lungs were dissected free of nonpulmonary tissue, weighed, and then dried to a constant weight in an oven at 130° C. The lung wet/dry weight ratio was obtained by dividing the wet weight by the final dry weight.

Histopathlogic studies of the lungs. Histopathologic examination of the lungs was performed 6 h after administration of LPS. Samples were fixed with 10% formalin, embedded in paraffin, sectioned into 6-μm pieces, and stained with hematoxylin-eosin. Samples were analyzed by a pathologist blinded to the animals' group assignment. Ten randomly selected fields per slide were read under oil at magnification ×1,000 by a pathologist. Fields containing large vessels or bronchi were excluded. The number of neutrophils per field was counted and normalized to the number of alveoli per field to control for lung infiltration.

Measurement of the plasma level of TNF-α in rats. Saline or cepharanthine (10 mg/kg) was administered intraperitoneally 30 min before an intravenous injection of LPS (5 mg/kg). Plasma samples were obtained 90 min after injection of LPS. Plasma levels of TNF-α were determined using an enzyme-linked immunosorbent assay (ELISA) kit for rat TNF-α (Genzyme, Cambridge, MA).

Isolation and cultivation of human monocytes. Peripheral blood mononuclear cells were isolated from buffy coats obtained from healthy volunteer blood donors by centrifugation on Nyco Prep 1.077 (Nycomed Pharma AS, Oslo, Norway) (13). The medium and buffer solutions were monitored for contamination with LPS using the Endotoxin Test-D (Seikagaku-kogyo, Tokyo, Japan). Mononuclear cells were cultured in RPMI 1640 (Gibco, Grand Island, NY) plus 10% fetal calf serum (Gibco) and then incubated in plastic dishes (Falcon 1058; Becton Dickinson, Lincoln Park, NJ) for 2 h at 37° C in a humidified 5% CO₂ incubator. Lymphocytes were removed from adherent monocytes by repeated rinsing with serum-free RPMI 1640. Staining with Turk's solution and for nonspecific esterase activity confirmed that > 90% of harvested cells were monocytes. Cells were adjusted to a volume of 5.0 × 10⁵/ml in RPMI 1640 and then stimulated with LPS (20 ng/ml) for 16 h at 37° C in a humidified 5% CO₂ incubator in the presence or absence of various concentrations of cepharanthine. After incubation, cell suspensions were centrifuged at 10,000 rpm for 10 min in an Eppendorf microcentrifuge (Model 5412; Hamburg, Germany). Levels of TNF-α and interleukin-1β (IL-1β) in supernatant fractions were determined using an ELISA kit for human (Otsuka Pharm. Co., Tokyo, Japan).

Preparation of neutrophils from normal human blood. Heparinized venous blood obtained from normal volunteers was mixed with an equal volume of 2% dextran solution and allowed to stand for 30 min to permit erythrocyte sedimentation. The supernatant was centrifuged and the precipitate fraction was collected. Neutrophils were isolated by a Nyco Prep 1.077. Contaminated erythrocytes were removed by hemolysis with 0.2% saline for 25 s. The resulting preparation, which contained more than 95% neutrophils, was washed twice with phosphate-buffered saline. Cell viability of 95% or higher was confirmed by the trypan blue dye exclusion test. Cells were suspended in phosphate-buffered saline in a volume of 5,000/μl.

Neutrophil elastase release. The neutrophil suspension (5,000/μl) in phosphate-buffered saline was mixed with 5 μg/ml of cytochalasin B to prime the cells, stimulated with 5 μg/ml of fMLP in the presence or absence of cepharanthine (14). After incubation for 30 min at 37° C, neutrophil suspensions were centrifuged at 5,000 × g for 10 min at 4° C. Neutrophil elastase activity in supernatants was measured using a chromogenic substrate (S-2484; Chromogenix AB, Stockholm, Sweden), according to a previously described method (15).

O₂ ⁻ production by neutrophils. Neutrophil production of O₂ ⁻ was measured by a chemiluminescence assay using a luminescence reader (type BLR-201; Aloka Co., Tokyo, Japan), as previously described (16). In brief, neutrophil suspensions (1.0 ml) were mixed with 10 μg/ ml of luminol dissolved in physiological saline and various concentrations of cepharanthine for 5 min at 37° C. The mixture was stimulated with opsonized zymosan (1.0 mg/ml) and changes in chemiluminescence activity were monitored continuously. The peak chemiluminescence intensity was determined by subtracting the basal chemiluminescence intensity (chemiluminescence activity before the addition of opsonized zymosan) from the maximal chemiluminescence intensity (maximal count/minute).

Neutrophil aggregation. Neutrophil aggregation was determined with an electric whole blood aggregometer (Chrono-Log, Havertown, PA) using a modification of a previously described method (17). In brief, 100 μl of the neutrophil suspension (5,000/μl) in phosphate-buffered saline was mixed with 900 μl of modified Tyrode's solution (NaCl, 8.0 g/L; MgCl₂ 6 H₂O, 0.427 g/L; CaCl₂, 0.2 g/L; KCl, 0.2 g/L; d-glucose, 1.0 g/L; NaHCO₃, 1.0 g/L; NaH₂PO₄ 2 H₂O, 0.065 g/L) with or without cepharanthine and incubated for 5 min at 37° C. The cell suspension was then stimulated with 2 × 10⁻⁸ M fMLP. Neutrophil aggregation was monitored for 20 min with the aggregometer. Aggregation is expressed in terms of changes in electrical impedance (17).

Intracellular ionized calcium concentration ([Ca²⁺]i) was measured as previously described (18). Briefly, neutrophils isolated as described above were suspended at 5 to 10 × 10⁶/ml in RPMI 1640 with 10% fetal calf serum and 2.5 μg/ml Indo-1 acetoxymethyl ester (Dojindo Laboratories, Kumamoto, Japan) for 30 min at 37° C. Cells were then washed twice and resuspended at 1 × 10⁶/ml in RPMI 1640 with 10% fetal calf serum. Immediately before analysis of [Ca²⁺]i, cells were washed in buffer A (140 mM NaCl, 3 mM KCl, 1 mM MgCl₂, 10 mM glucose, 1 mM CaCl₂, and 20 mM Hepes at pH 7.23) and suspended (1.0 × 10⁶ cells in a volume of 1.7 ml) in a thermostatically controlled (37° C) cuvette. Fluorescence emission was measured in a spectrophotometer (Hitachi 850; Hitachi Ltd., Tokyo, Japan) using an excitation wavelength of 331 nm and an emission wavelength of 410 nm. After equilibration of fluorescence to a stable baseline, the cells (1.0 × 10⁶) were stimulated with fMLP (0.3 μM), and fluorescence assessment was continued. Fluorescence levels were calibrated in terms of [Ca²⁺]i after each experiment by lysing the cells with 0.1 to 0.2% Triton X-100, and measuring fluorescence in the presence (Fmax: buffer A containing 1 mM Ca²⁺) and the absence (Fmin: buffer A containing 4 mM EDTA) of calcium. [Ca²⁺]i was calculated from the following formula: [Ca²⁺]i = Kd(F-Fmin)/(Fmax-F), where Kd = 180 nM for Indo-1 at pH 7.2 (19) and F is the fluorescence of the unknown.

Values are expressed as the mean ± SD. Data were analyzed by analysis of variance and Scheffé's post hoc test for group pairs for multiple comparisons, and by the unpaired t test for single comparison. Statistical significance was defined as p < 0.05.