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J Thorac Cardiovasc Surg 2001;122:123-128
© 2001 The American Association for Thoracic Surgery
Cardiopulmonary Support and Physiology (CSP) |
From the British Heart Foundation, Cardiac Surgerya and Cardiovascular Medicineb Units, Hammersmith Hospital, National Heart and Lung Institute, Imperial College School of Medicine, London, United Kingdom.
Received for publication Aug 30, 2000. Revisions requested Nov 14, 2000; revisions received Dec 4, 2000. Accepted for publication Jan 17, 2001. Address for reprints: R. Clive Landis, PhD, BHF Cardiovascular Medicine Unit at Hammersmith Hospital, National Heart and Lung Institute, Imperial College School of Medicine, Du Cane Rd, London W12 0NN, United Kingdom (E-mail: r.landis{at}ic.ac.uk).
Abstract
Background: Cardiopulmonary bypass surgery is often accompanied by a systemic inflammatory response, which can lead to postoperative complications in high-risk patients. This is mediated in part through a systemic rise in inflammatory cytokine levels and the sequestration of leukocytes within organs. Aprotinin has previously been shown to exert an anti-inflammatory effect by preventing the capacity of leukocytes to transmigrate through vascular endothelium. Here we have focused on whether aprotinin has an effect on endothelial cell activation and adhesion molecule expression in response to tumor necrosis factor–
, particularly with reference to whether aprotinin inhibits tumor necrosis factor–stimulated neutrophil transendothelial migration.
Methods and Results: Intercellular adhesion molecule–1, vascular cell adhesion molecule–1, and E-selectin expression was studied in tumor necrosis factor––activated human umbilical vein endothelial cells in the presence of aprotinin at 200, 800, and 1600 kIU/mL. Aprotinin inhibited tumor necrosis factor––stimulated expression of intercellular adhesion molecule–1 (P = .019 at 1600 kIU/mL) and vascular cell adhesion molecule–1 (P = .003 at 1600 kIU/mL) but not E-selectin. Similar results were obtained in the dermal microvascular endothelial cell line, HMEC-1, which exhibited diminished intercellular adhesion molecule–1 expression in the presence of aprotinin (P = .040 at 800 kIU/mL and P < .001 at 1600 kIU/mL). Aprotinin also significantly inhibited neutrophil transmigration across tumor necrosis factor––activated human umbilical vein endothelial cells (P = .046 at 1600 kIU/mL).
Conclusions: We have demonstrated that aprotinin inhibits intercellular adhesion molecule–1 and vascular cell adhesion molecule–1, but not E-selectin, expression on tumor necrosis factor––activated endothelial cells and that transendothelial migration by neutrophils is also specifically suppressed under these conditions. Our results indicate that endothelial cells can be specifically targeted by aprotinin, therefore adding to our understanding of the anti-inflammatory mechanism of action of aprotinin during cardiopulmonary bypass.
Cardiopulmonary bypass (CPB) surgery is frequently accompanied by a systemic inflammatory response, which can cause postoperative complications, lengthened duration of hospital stay, and, in the worst case scenario, multiple organ failure.
1-4 The etiology of this inflammatory response has been traced to the stress of surgery and contact activation of platelets and leukocytes within the bypass circuit, which leads to an increase in circulating cytokine levels such as tumor necrosis factor (TNF)–, interleukin-1, interleukin-6, and interleukin-8. 5-7 Inflammatory cytokines in turn cause endothelial cell (EC) activation and expression of vascular adhesion molecules involved in recruitment of leukocytes to sites of inflammation or tissue injury. Acute organ injury, which is most often observed in the ischemic lung, results from the accumulation of neutrophils within the affected organ and the accompanying release of histotoxic mediators from infiltrated cells. 8
Leukocyte emigration from the vasculature is governed by an orderly series of contact events between leukocytes and endothelium, involving adhesion molecules of the selectin, integrin, and immunoglobulin superfamilies. 9 In general, the initial leukocyte tethering and rolling event is mediated via the selectins E-, L-, and P-selectin, which are specialized to carry out transient adhesive contacts under conditions of hydrodynamic shear. The subsequent firm adhesion step is mediated by the leukocyte integrins lymphocyte function–associated antigen-1, macrophage antigen-l, and very late antigen-4, which recognize immunoglobulin superfamily counter-ligands on the endothelial surface, including intercellular adhesion molecule (ICAM)–1, ICAM-2, and vascular cell adhesion molecule (VCAM)–l. Least is known about the final extravasation step, which involves contributions from the immunoglobulin superfamily molecules ICAM-1, ICAM-2, and platelet EC adhesion molecule–1, as well as leukocyte-derived proteases that digest endothelial junctions and matrix barriers ahead of the migrating cell. 10
Although anti-inflammatory effects of aprotinin in CPB have been demonstrated in clinical trials, 11-13 animal models of acute lung injury, 14,15 and in vitro models of neutrophil activation, 16,17 the action of aprotinin on the leukocyte-endothelial cascade remains poorly understood. We have previously used intravital microscopy to study the effect of aprotinin on leukocyte recruitment in response to a topical chemoattractant, N-formyl-methyl-leucyl-phenylalanine, within the rat mesenteric microcirculation. These experiments showed that aprotinin inhibited the extravasation step of the leukocyte-EC adhesion cascade but had no effect on the rolling or firm adhesion steps. 18 Parallel in vitro experiments supported these observations by showing that aprotinin dose-dependently inhibited neutrophil transmigration through human umbilical vein endothelial cell (HUVEC) monolayers in response to the chemoattractants N-formyl-methyl-leucyl-phenylalanine, interleukin-8, or platelet-activating factor. 18 Although this study was seen as relevant to the sequestration of neutrophils during acute lung injury, 19,20 no in vitro model system can adequately model the complexities of the in vivo situation, and in the present study the emphasis has been placed on a more endothelial cell–specific agonist, TNF-. 21
TNF- promotes leukocyte–EC interactions by upregulating expression of a number of EC adhesion molecules including E-selectin, ICAM-1, and VCAM-1. 21 Here we have focused on whether aprotinin inhibits the ability of TNF to activate ECs, and, if so, whether this influences leukocyte-EC interactions.
Methods
Reagents and antibodies
Aprotinin, dexamethasone, gelatin, and fibronectin were purchased from Sigma-Aldrich Chemical Co Ltd (Poole, Dorset, United Kingdom). TNF- was the gift of Dr Martyn Robinson, Celltech Chiroscience Ltd (Slough, United Kingdom). Anti-ICAM-1 monoclonal antibody 15.2 was the gift of Dr Nancy Hogg, Imperial Cancer Research Fund (London, United Kingdom). The anti-E-selectin monoclonal antibody 1.2B6 and anti-VCAM-1 monoclonal antibody 1.4C3 were generated within our group as previously described. 22
EC culture
HUVECs were obtained from umbilical cords by collagenase type II (Boehringer-Mannhein, Lewes, Sussex, United Kingdom) digestion, as previously described. 22 Human umbilical cords were cleaned with 70% ethanol, and the umbilical vein was identified and flushed with Hanks balanced salt solution to remove blood. The vein was then filled with 10 mL of 0.1% collagenase (Boehringer) in Hanks balanced salt solution, the vessel ends clamped, and the cord incubated for 10 minutes at 37°C. The collagenase digest was retained, and the cord was flushed through with 20 mL of Hanks balanced salt solution to collect the remaining HUVECs. The ECs were pelleted by centrifugation at 200g for 5 minutes and resuspended in HUVEC growth medium consisting of medium M199 supplemented with 20% heat-inactivated (56°C for 30 minutes) fetal calf serum (Hyclone Laboratories Inc, Logan, Utah), 100 IU/mL penicillin, 100 µg/mL streptomycin, 2 mmol/L L-glutamine, 10 U/mL heparin, and 30 µg/mL EC growth factor (Sigma) before plating out on 1% v/v gelatin coated 25-cm2 tissue culture flasks. Once confluent, cells were passaged with trypsin (0.5 mg/mL)–ethylenediamine tetraacetic acid (0.2 mg/mL) for 1 minute at 37°C followed by inactivation with cold phosphate-buffered saline solution and fetal calf serum. Cells were sedimented at 210g for 10 minutes at 4°C, resuspended in medium, and transferred into gelatin-coated 75-cm2 tissue culture flasks, having been split in a 1:2 or 1:3 ratio. The HUVECs were passaged twice weekly and used between passages 2 and 5.
The human dermal microvascular EC line HMEC-1 (the gift of Dr Edwin Ades, Center for Disease Control, Atlanta, Ga) was maintained in culture medium MCDB 131 (Life Technologies, Paisley, United Kingdom), grown to confluence, and passaged as described above for HUVECs.
Neutrophil isolation from human venous blood
The technique used for isolation of human neutrophils has been previously described. 18 Peripheral venous blood from 5 healthy nonsmoking human donors was anticoagulated immediately by 2 mL of sodium citrate 3.8% (Pharma Hameln, Hameln, Germany) per 25 mL of blood and aliquoted into polypropylene tubes. After centrifugation at 3000 rpm for 10 minutes the plasma was removed, and cells were mixed with 6 mL of 6% high molecular weight dextran (Dextran T500; Amersham-Pharmacia Biotech, Amersham, United Kingdom) and 20 mL phosphate-buffered saline solution. After 25 minutes to sediment red cells, the leukocyte-rich upper layer was harvested, centrifuged at 1200 rpm for 5 minutes, resuspended in 2 mL of autologous plasma, and layered onto a 2-step (74% and 50%) Percoll (Amersham-Pharmacia) gradient. After centrifugation at 1300 rpm for 10 minutes, neutrophils were collected from the top of the 74% interface, washed twice in RPMI 1640 medium containing 2% fetal calf serum, and counted.
Neutrophil transmigration experiments
For transmigration experiments, HUVECs were coated onto 3.0-µm pore-size cell culture inserts (Falcon; Becton-Dickinson, Franklin Lakes, NJ) as previously described. 18 One day before transmigration, cell culture inserts were coated with 100 µL of phosphate-buffered saline solution containing 50 µg/mL fibronectin for 2 hours. After excess fibronectin had been rinsed off, inserts were placed into 24-well plates (Nunclon; Nalge Nunc International, Roskilde, Denmark), and 1 x 105 HUVECs were added to each insert. After overnight incubation, ECs were preincubated for 60 minutes with aprotinin at 200 kIU/mL, 800 kIU/mL or 1600 kIU/mL, or medium. So that the possible effects of aprotinin on neutrophils could be avoided, aprotinin was washed out from HUVEC lawns with 2 rinses of RPMI 1640, and, subsequently, RPMI 1640 containing TNF- was added at 2 ng/mL to the lower chamber. Four and one-half hours after administration of TNF-, neutrophils were added at 1 x 106 to the upper surface of filter inserts. Neutrophil transmigration was measured at 60 minutes by counting cells that had migrated into the lower chamber. Each experiment was performed in duplicate and repeated 4 times with different HUVEC and neutrophil isolates, no two isolates being alike.
Flow cytometric analysis of ICAM-1, VCAM-1, and E-selectin expression
HUVECs or HMEC-1 were isolated and cultured in petri dishes. Cells were used between passages 2 and 5. At confluence, ECs were preincubated with aprotinin at 200 kIU/mL or 1600 kIU/mL for 60 minutes before challenge with TNF- at 2 ng/mL. After 4.5 hours of culture in the presence of TNF-, cells were harvested with trypsin (0.5 mg/mL)-ethylene diaminetetraacetic acid (0.2 mg/mL), resuspended in medium, transferred into polypropylene tubes, and incubated with primary antibodies 15.2, 1.4C3, and 1.2B6 for 15 minutes on ice. An isotype-matched antibody was used as a control. Preliminary experiments demonstrated that a time period of 4.5 hours was necessary to achieve significant simultaneous TNF-–induced up-regulation of ICAM-1, VCAM-1, and E-selectin. After 3 washes with phosphate-buffered saline solution, fluorescein isothiocyanate coupled goat anti-mouse immunoglobulin G secondary antibody was added at the manufacturer's recommended concentration (Sigma Chemical Co, Dorset, United Kingdom), and incubation continued for an additional 15 minutes on ice. After 3 additional washes in phosphate-buffered saline solution, flow cytometric analysis was carried out with an EPICS XL flow cytometer (Coulter Electronics Ltd, Luton, United Kingdom). The staining intensity of each test antibody was initially corrected with respect to an isotype-matched control antibody to obtain a relative staining intensity (RFI). To average RFIs between experiments, it was necessary to normalize expression relative to the control sample (in the absence of any agonists) to correct for the natural variability in adhesion molecule expression between fresh EC isolates. Final results were expressed as fold increase over control.
Statistics
The effect of aprotinin (200, 800, and 1600 kIU/mL) on TNF-–induced adhesion molecule expression and neutrophil transmigration was analyzed with a 1-way analysis of variance in conjunction with a Student-Neuman-Keuls post-test (SPSS Inc, Chicago, Ill). The effect of 10–5 mol/L dexamethasone on TNF-induced adhesion molecule expression was analyzed with a paired t test (SPSS Inc).
Results
Aprotinin inhibits ICAM-1 and VCAM-1, but not E-selectin, expression on HUVECs
Figure 1 shows the effect of aprotinin on the TNF-–induced expression of ICAM-1, VCAM-1, and E-selectin. All 3 adhesion molecules were significantly up-regulated on HUVECs by 4.5-hour culture in the presence of 2 ng/mL TNF-. Aprotinin dose-dependently inhibited expression of ICAM-1 (P = .019 at 1600 kIU/mL)(Figure 1, A) and VCAM-1 (P = .003 at 1600 kIU/mL)(Figure 1, B), but not E-selectin(Figure 1, C). As a control, dexamethasone inhibited TNF-–induced expression of all 3 adhesion molecules, consistent with previous reports of its inhibitory action against NF-
B agonists. 23 ICAM-1 expression in the presence of 10–5 mol/L dexamethasone was 60.18% ± 21.53% (P = .026) relative to TNF only, VCAM-1 was 64.35% ± 23.81% (P = .011), and E-selectin was 68.06% ± 21.86% (P = .020).
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–induced ICAM-1 expression in HMEC-1 (P = .040 at 800 kIU/mL and P < .001 at 1600 kIU/mL).
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–activated HUVECs–activated lawns of HUVECs grown onto Transwell filter inserts (3-µm pore size) in the presence and absence of aprotinin. Figure 3 shows that exposure of HUVECs to TNF- for 4.5 hours resulted in an up-regulation of neutrophil transmigration, but this was significantly reduced by pretreatment of the endothelial lawn for 60 minutes with aprotinin (P = .046 at 1600 kIU/mL).
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We have focused on identifying an endothelial component to the protective action of aprotinin during CPB-related inflammation. Our results show that aprotinin blunts TNF-induced ICAM-1 expression in ECs of both large vessel and dermal microvascular origin. Furthermore, although TNF-–induced ICAM-1 and VCAM-1 levels are both inhibited by aprotinin in HUVECs, E-selectin expression is not. The possible functional relevance of these findings is indicated by the ability of aprotinin to inhibit neutrophil transmigration across TNF-–activated HUVECs.
The decrease in neutrophil trafficking may be due to reduced ICAM-1 expression, because neutrophils do not use the very late antigen-4:VCAM-1 axis but rely almost entirely on binding of ICAM-1 to lymphocyte function–associated antigen-1 or macrophage antigen-1 to carry out firm adhesion and extravasation. 24 The firm adhesion step is apparently not targeted by aprotinin, because neutrophil adhesion, with the use of a parallel plate flow-chamber, remains unchanged in the presence of aprotinin (data not shown), consistent with previous in vivo work. 18 It is unlikely that proteases of a neutrophil origin were inadvertently targeted, because aprotinin was carefully washed away from endothelial monolayers before the addition of neutrophils. This washing step may also explain the requirement for a relatively high dose of aprotinin (1600 kIU/mL) to achieve significant inhibition of transmigration compared with levels used in the clinic (200 kIU/mL). Extrapolation from the in vitro to the clinical situation is difficult, however, because previous studies have shown that although high doses of aprotinin (800 and 1600 kIU/mL) are required to inhibit neutrophil transmigration in vitro, much lower levels (<200 kIU/mL) are required to achieve significant inhibition in vivo. 18
It is interesting to note that aprotinin specifically inhibited expression of the 2 immunoglobulin superfamily molecules, ICAM-1 and VCAM-1, but not the selectin adhesion molecule E-selectin. This was a surprise, because all 3 genes are known to be induced by similar NF-B–dependent signaling pathways and all are inhibitable by the administration of dexamethasone. Our results suggest that a protease-sensitive pathway may play a necessary role in optimal ICAM-1 and VCAM-1 expression. Obvious candidate signaling receptors are the protease-activated receptor (PAR) family, 25 which includes the classic thrombin receptor, PAR1, that we have previously shown is blocked by aprotinin on platelets. 26 ECs express PAR1 and PAR2, which are activated by proteolytic cleavage with thrombin and trypsin, respectively, 27 and because aprotinin is known to block these in vitro, 28,29 it is possible that PAR activation in HUVECs may contribute to the expression of ICAM-1 and VCAM-1. This possibility is supported by the fact that overt stimulation through thrombin can up-regulate ICAM-1 and VCAM-1 expression. 30 The PARs are not strictly ligand-specific and can be activated by a wide variety of serine proteases, therefore leaving open the possibility that an unknown protease of endothelial origin may be targeted by aprotinin. Growing evidence that the PARs can be cofactored by endothelial surface molecules in -cis further complicates the picture but enhances the likelihood that these receptors are involved, either directly or indirectly, in mediating the effects reported here with aprotinin. 31
The results presented here have added an endothelum-specific dimension to the increasingly complex mechanism of anti-inflammatory action of aprotinin in relation to CPB. The wide distribution of protease targets within the coagulation and fibrinolysis cascades, as well as the recent recognition that cell-associated PARs may be targeted by aprotinin, 26 may explain how aprotinin can achieve such diverse anti-inflammatory effects at the level of (1) cytokine synthesis, (2) contact activation of platelets and neutrophils, (3) EC activation, and (4) leukocyte extravasation. Although it will be important to evaluate future generations of protease inhibitors with higher specific activity or longer half-life than aprotinin, the attraction of aprotinin remains that as a nonspecific serine protease inhibitor it can target a broad range of substrates both in the soluble phase and associated with the cell surface, which may prove beneficial in the management of hemostasis and inflammation in CPB surgery.
References
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