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J Thorac Cardiovasc Surg 2002;124:321-332
© 2002 The American Association for Thoracic Surgery
Surgery for Acquired Cardiovascular Disease (ACD) |
From the Department of Medical Sciences, Clinical Chemistry, Laboratory for Coagulation Research,a the Departments of Oncology, Radiology, and Clinical Immunology,b and the Department of Cardiothoracic Surgery,c University Hospital, Uppsala, Sweden.
This study was supported by the Foundation for Strategic Research in Sweden and the Medical Research Council.
Received for publication Aug 21, 2001. Revisions requested Oct 16, 2001; revisions received Dec 26, 2001. Accepted for publication Dec 28, 2001. Address for reprints: Agneta Siegbahn, MD, PhD, Laboratory for Coagulation Research, Clinical Chemistry, Department of Medical Sciences, University Hospital, SE-751 85 Uppsala, Sweden (E-mail: Agneta.Siegbahn{at}clm.uas.lul.se).
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Abstract |
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Introduction |
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1 Thrombin generation during cardiac surgery with CPB might be caused by the surface of the extracorporeal unit, with the oxygenator, pump, and connections between the patient and the extracorporeal device. 2 The surgical trauma with exposure of tissue factor (TF), the principal initiator of blood coagulation, is also suggested to be of major importance for thrombin generation. 3-5 Previously, it was thought that the activation of coagulation during CPB occurred mainly through contact activation. However, current evidence suggests that activation of the TF-induced pathway might be the most important mechanism of thrombin generation during this type of operation. 6-8 Heparinization is required to prevent clotting in the extracorporeal circuit during cardiac surgery, and heparin coating has been reported to improve the biocompatibility of artificial surfaces in contact with circulating blood. 9 Moreover, the enhanced biocompatibility of CPB circuits by means of heparin coating has led to the idea of reducing the systemic heparinization during cardiac surgery. This was shown to be clinically safe, with a lower incidence of homologous transfusion and thromboembolic complications, although no differences in markers for inflammation, coagulation, and fibrinolysis. 10-12
Retransfusion of shed mediastinal blood (SMB) was introduced by Schaff and colleagues 13 in 1978 and has become a widely used method during and after cardiac surgery. 14,15 However, a number of studies have shown that the composition of the shed blood is far from normal. 16,17 SMB after CABG is characterized by extraordinary activated coagulation and fibrinolysis. 18-20 Furthermore, high levels of proinflammatory cytokines are found in shed blood. 20 However, the interpretations of the outcome of retransfusion of this blood differs; some studies have concluded that retransfusion of shed blood deteriorates hemostasis, fibrinolysis, and inflammation, whereas others suggest that it does not. 16,17,20-22
A new heparin surface, the Corline Heparin Surface (CHS), has been prepared by the use of a macromolecular conjugate of covalently bonded heparin. In this study we have, for the first time, investigated the effects in CPB of the CHS in combination with different concentrations of systemic heparin on leukocytes, platelets, coagulation, and inflammatory responses compared with effects achieved with an uncoated device. The aim was to further study the combination of cell and tissue activation, inflammation, and coagulation in both systemic and shed blood during and after cardiac surgery with CPB.
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Material and methods |
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The study was planned and performed in 2 parts, with a short delay between the 2 parts giving time for analyses of some important parameters to survey the in vivo effects of the new surface. In the first part patients were randomly assigned to group 1 (n = 15) for the use of an uncoated circuit (Jostra Medizintechnic AG, Hirrlingen, Germany) and to group 2 (n = 15) for a completely heparin-coated (Corline Heparin Surface; Corline Systems AB, Uppsala, Sweden) but otherwise identical device. Both groups received a standard dose of systemic heparin, corresponding to an activated clotting time (ACT) of 480 seconds. In the second part of the study, the CHS was used in all patients (groups 3 and 4). In group 3 (n = 15) patients were randomized to a low dose of systemic heparin (ACT of 300 seconds) and group 4 (n = 15) to a high dose of systemic heparin (ACT of 600 seconds).
No autotransfusion of SMB was performed during this study.
Heparin surface
The CHS was applied to a complete set of an extracorporeal circuit (tubing, cannula, oxygenator, and reservoir). The CHS uses a unique macromolecular conjugate in which approximately 70 heparin molecules are covalently linked to a polymer carrier having a molecular weight of 50 kd. The specificity of the covalent linkages ensures that the antithrombin III-binding pentasaccharide sequence of heparin are left intact. The CHS coating comprises a conditioning layer of a polymeric cationic amine onto which the macromolecular conjugate is irreversibly attached by means of multiple ionic interactions. The surface concentration of heparin is 0.5 to 1 µg/cm2, and the capacity to bind antithrombin is 2 to 4 pmol/cm2. 23 An extracorporeal closed-loop system modified with CHS used during extracorporeal circulation for 24 hours in pigs without any systemic dose of heparin had no effect on the clotting time (data submitted to the Swedish Medical Products Agency).
Anesthesia and analgesia
Patients were premedicated with morphine or morphine-scopolamine on the morning of the operation. Anesthesia followed conventional procedures with thiopental sodium used for induction and a combination of pancuronium bromide and inhaled isoflurane used during the operation. After weaning from CPB, propofol in continuous infusion was used as an anesthetic agent. Alfentanil hydrochloride and fentanyl citrate were used for intraoperative analgesia.
Cardiopulmonary bypass
Techniques and equipment for CPB were similar in all patients. The standard set consisted of a Stöckert roller pump (Stöckert Instrumente GMBH, Munich, Germany) with a Quadrox hollow-fiber oxygenator (Jostra) and a hard-shell venous reservoir (Jostra). The tubing, including cannula, was CHS coated from tip to tip (9 patients had uncoated cannulas because of surgeons' preferences). The circuit was primed with 1500 to 2000 mL of asanguineous fluid containing 5000 IU of heparin. The operation was performed with moderate hypothermia (32°C-35°C), and the patients were rewarmed to a rectal temperature of 36°C before weaning from CPB. The flow was nonpulsatile, initially 2.4 L · m-2 · min-1, and gradually reduced to 2.0 L · m-2 · min-1 during hypothermia.
Anticoagulation and operative procedures
Patients in groups 1 and 2 were anticoagulated by means of intravenous administration of heparin administered as a bolus of 300 IU/kg body weight after completed dissection of the internal thoracic artery and before cannulation for CPB. In groups 3 and 4 the bolus doses of heparin were 200 and 400 IU/kg body weight, respectively. If necessary, additional doses of heparin were given to maintain the desired ACT. After decannulation, heparin was reversed with protamine chloride in a 1:1 proportion to the administrated heparin dose. ACT was measured repeatedly throughout the operation with an ACT II (Medtronic Inc, Minneapolis, Minn) to assess adequate anticoagulation. Cardioplegic arrest was achieved with antegrade infusion of modified St Thomas' cardioplegic solution at 4°C through the aortic root. During the CPB procedure, cardiotomy suction was used. Blood remaining in the CPB circuit after decannulation was collected in an infusion bag and immediately retransfused. Postoperatively, shed blood was collected in the reservoir, but no retransfusion was performed.
Blood sampling and laboratory procedures
Blood samples were drawn from a catheter in the radial artery after induction of anesthesia, at sternotomy, at the start and end of CPB, 30 minutes after protamine administration, and 3, 8, and 20 hours after CPB. Sampling was also performed from the pericardial cavity at the end of CPB and from the drainage reservoir (after gentle mixing of the reservoir) 3 hours after CPB. Vacutainer tubes containing Na-citrate or ethylenediamine tetraacetic acid K3 (Becton Dickinson, Meylan Cedex, France) were used. One citrated sample was immediately brought to the laboratory for cell count and analysis of cellular surface antigens by means of flow cytometry. The other samples were immediately centrifuged at 2000g for 20 minutes, and plasma was frozen and stored in aliquots at -70°C until analysis.
Platelet and leukocyte differential counts were determined with an automatic cell counter (Coulter STKS; Coulter Electronics, Hialeah, Fla). Plasma levels of prothrombin fragment 1+2 (F1+2), thrombin-antithrombin (TAT), plasmin/
2-antiplasmin complex (PAP), human myeloperoxidase (MPO), and the cytokines interleukin (IL) 6, IL-8, IL-10, and soluble IL-2 receptor (sIL-2R) were quantified by means of sandwich enzyme-linked immunosorbent assay techniques (Enzygnost for F1+2, TAT, and PAP from Behringswerke, Marburg, Germany; Bioxytech for MPO enzyme immunoassay from Oxis International, Inc Portland, Ore; and Quantikine for IL-6, IL-8, IL-10 and sIL-2R from R&D Systems, Abingdon, United Kingdom). In a reference group of 71 healthy individuals, reference intervals (2.5th-97.5th percentile) for F1+2 and TAT were 0.4 to 1.5 nmol/L and 1.2 to 5.0 µg/L, respectively, with respective medians of 0.8 nmol/L and 2.3 µg/L. For both assays, the intra-assay and interassay variations were 5% and 9%. In a reference group of 466 healthy individuals, reference intervals (2.5th-97.5th percentile) for PAP was 120 to 700 µg/L, with a median of 290 µg/L. The intra-assay and interassay variations were 4% and 7%, respectively. Plasma soluble fibrin (sF) was analyzed with an assay using a chromogenic, plasmin-specific substrate (Bericrom FM, Behring Diagnostics GmbH). In 73 healthy individuals aged 21 to 66 years (median, 28 years), the reference interval (2.5th-97.5th percentile) for sF was 17 mg/L or less (median, 9.0 mg/L; range, 3.2-16.6 mg/L), and the intra-assay and interassay variations had a total coefficient of variation of 7.9%. Reference intervals for the cytokines were according to the manufacturer:
The intra- and interassay variations were 1.7% and 3.3% for IL-6, 5.4% and 9.7% for IL-8, 4.3% and 7.5% for IL-10, and 6.1% and 6.0% for sIL-2R, respectively.
Determination of activated coagulation factor XI-antithrombin complex (FXIa-AT), activated coagulation factor XII-antithrombin complex (FXIIa-AT), and activated coagulation factor XII-C1-esterase inhibitor complex (FXIIa-C1 INH) were performed by using a solid-phase enzyme-linked immunosorbent assay according to the method of Sanchez and colleagues. 24 The intra-assay coefficient of variation was less than 4% in all assays. Cross-reactions between antibodies to FXIa and FXIIa or antibodies against AT and C1 INH were less than 5%.
No correction for hemodilution was made.
Flow cytometry
Leukocytes
Surface antigen expression was analyzed by using a Coulter Epics XL-MCL flow cytometer (Beckman Coulter, Fullerton, Calif). Whole blood was labeled with a murine monoclonal flourescein isothiocyanate (FITC)-conjugated anti-TF antibody (TF4508-CJ; American Diagnostica, Greenwich, Conn) or an anti-CD11b antibody (Dakopatts A/S, Glostrup, Denmark). A murine FITC-conjugated irrelevant antibody of the same subtype was used as a negative control. The samples were incubated on ice for 30 minutes and lysed with Lyse (Beckman Coulter), washed, and suspended in PBS. Flow cytometric analysis was then performed. Gating of the cells was carried out with forward- and side-scatter parameters, and the percentage of positive cells and mean fluorescence intensity were determined.
Platelets
Platelet-rich plasma was isolated by means of centrifugation at 140g for 10 minutes at room temperature. Platelet-rich plasma was added to tubes containing HEPES buffer and FITC-labeled chicken polyclonal anti-P-selectin antibody (Immunsystem AB, Uppsala, Sweden) or FITC-labeled chicken polyclonal anti-whole-platelet antibody (Biopool AB, Umeå, Sweden). The samples were incubated in duplicate for 10 minutes at room temperature and were then diluted and fixed with ice-cold PBS containing 1% paraformaldehyde. No washing steps were used. For P-selectin analysis, gating of the cells in the flow cytometer was carried out by using forward- and side-scatter parameters, and for microparticle analysis, gating was performed by using forward scatter and fluorescence 1 (anti-whole-platelet antibodyFITC). Platelets with fluorescence of less than a preset cut-off level were identified as microparticles. The percentage of positive cells was determined.
Statistical analysis
Statistica for Windows (StatSoft, Tulsa, Okla) was used for all statistical analyses. The results are presented as medians and 25th-75th percentile ranges. The Friedman analysis of variance test was used for analysis of time series within each group. When significant differences were obtained, these were further evaluated by using the Wilcoxon matched pairs test for comparison between 2 samples within the series. The Kruskal-Wallis analysis of variance test was used for an overall comparison between the groups. When significant differences were obtained, these were further evaluated with the Mann-Whitney U test for intergroup comparison. No adjustment for multiple testing was done. Hence this study should be interpreted more as an exploratory, rather than a confirmative, study.
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).
Platelet activation
The number of platelets in circulating blood was decreased during the operation in all groups, from 160 x 109/L (134-182 x 109/L) to 107 x 109/L (77-136 x 109/L, P < .001). In the uncoated group, group 1, and the heparin-coated group with high systemic heparin, group 4, the decrease was even more pronounced in comparison with that in the heparin-coated group with standard heparin dose, group 2 (P = .03 and P = .02, respectively), probably because of the activation and adhesion in these groups. Forty-eight hours after the operation, the platelets had returned to presurgical levels in all groups. The expression of P-selectin on circulating platelets was increased during the operation in all groups (P < .001); however, this expression was less in group 2 than in group 1 (P = .004). This positive effect of the heparin surface was not affected by the systemic heparin dose (Figure 3). The formation of platelet microparticles in the circulating blood was not significantly altered in any group during the operation (data not shown).
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Discussion |
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11,25,26
Despite the fact that a number of experimental studies have demonstrated inhibition of FXa and thrombin as a result of immobilized heparin, many clinical studies using heparin-coated bypass circuits have failed to demonstrate reduced thrombin formation. 18,27 To our knowledge, only one study 28 with heparin coating (Duraflo II surface) showed reduced formation of thrombin. The seeming discrepancy between the experimental and clinical results with respect to activation of coagulation might reflect that a number of factors in addition to the artificial surfaces, such as pumping of blood, rheologic imperfections, surgical trauma, and retransfusion of pericardial and SMB come into play in the clinical setting.
The aim of the first part of this study was to investigate the biocompatibility of a newly developed heparin surface, the CHS. It should be emphasized that the CHS is a stable heparin coating with no demonstrable release of heparin and a high capacity to bind antithrombin. Judged by these characteristics, the CHS is comparable with the Carmeda Bioactive Surface but different from the Duraflo II, which is based on an ionic complex with sustained release of heparin and the cationic surfactant used in the complex with no demonstrable capacity to bind antithrombin.
Markers of inflammation, coagulation, and fibrinolysis were included in the analysis to obtain an overall view of processes activated at blood contact with biomaterials. The CHS significantly reduced the activation of coagulation, fibrinolysis, platelets, and inflammation in comparison with that on the uncoated surface in combination with a standard dose of systemic heparin. These results indicate that the CHS is more biocompatible than an uncoated surface, not only in terms of inflammatory parameters but also in terms of reduced thrombin generation.
The improved biocompatibility achieved by coating the surface of the extracorporeal device with heparin has initiated studies in which the dose of systemic heparin has been reduced. However, these studies have presented conflicting results. Some have shown a reduction of granulocyte activation 11 and a lower incidence of homologous transfusions, 12 whereas others found it clinically safe but not in favor to standard heparin levels in combination with available heparin surfaces. 10,29 Yet others have concluded that a reduction of systemic heparin should not be made because of an increased formation of thrombin. 30 An important difference between various clinical studies is the use of different surfaces and heparin doses at the same time. 30,31 This design makes it hard to distinguish between the effects of heparin coating and those induced by means of reduced anticoagulation. Thus in the second part of the present study, we investigated the effect of the CHS in combination with a reduced or increased systemic heparin dose compared with the standard systemic heparin dose used in part 1 in respect to inflammation, coagulation, and fibrinolysis.
In this study the reduced heparin dose was aimed at an ACT at 300 seconds, although this was not quite accomplished. The results were 400 seconds before and 343 seconds after CPB; however, these were significantly different from that seen in the coated group with a standard heparin level. The ratio between these 2 groups was constant during CPB (Table 1). The decrease of systemic heparin in combination with this new heparin-coated surface resulted in a slightly beneficial effect on inflammation reflected by the inflammatory marker MPO analyzed in this study. CD11b expression on the surface of leukocytes and data on coagulation and fibrinolysis, however, indicated elevated activation in the group with reduced heparin. These results might be explained by a requirement of a certain amount of soluble heparin to downregulate the procoagulant activity caused by the CPB and not accomplished by the surface per se. Heparin induces secretion of TF pathway inhibitor from endothelial cells in a dose-dependent manner, thus inhibiting TF/FVIIa. 32,33 Moreover, heparin can inhibit binding of fibrinogen, factor X, iC3b, and intercellular adhesion molecule 1 to Cd11b/CD18. 34 This might result in alteration of leukocyte functions involved in the activation of coagulation and inflammation.
The higher dose of systemic heparin (ACT of 600 seconds) in combination with the heparin surface was designed to clarify whether modulation of inflammation and coagulation by means of heparin could be further improved. However, the results from this group demonstrated the opposite, with increased activation of coagulation. Heparin is known to exert a variety of effects on cells in the vessel walls and circulation. Heparin potentiates platelet aggregation, recently shown to be caused by decreased nitric oxide production, an important endogenous inhibitor of platelet-mediated thrombosis. 35 This observation might in part explain the prothrombotic effects of the high dose of heparin.
Contact activation was reflected distinctly differently by the appearance of FXIIa-AT and FXIa-AT complexes, respectively. Whereas FXIa was reduced in group 2 compared with that seen in groups 3 and 4, there were no differences among the groups with respect to FXIIa. Quite interestingly, the generation of FXIIa started earlier, after sternotomy, with no further increase during CPB in groups 1 and 2. It has been reported that the capacity of the natural endothelium to inhibit FXIIa is impaired after systemic administration of heparin, 36 which might explain the early appearance of FXIIa, irrespective of the type of extracorporeal circuit, after heparinization of the patient. Because the generation of FXIa is greatly influenced by a thrombin feedback loop, 37 it seems plausible that the time course and relative concentrations of FXIa would coincide with those of thrombin formation. Reduced contact activation in connection with the use of Duraflo II has previously been reported. 7 However, this conclusion was based on reduced formation of kallekrein-CI INH complexes, with no direct measurements of FXIIA or FXIa or any signs of decreased thrombin formation.
TF is not normally expressed in the circulation. Monocytes in samples taken from the pericardial cavity have been shown to express approximately 2-fold increased TF on the cell surface 3 in a study performed with an uncoated extracorporeal device. Our results are confirming, with no differences between the groups. The use of cardiotomysuction in this study with recirculation of the blood might contribute to the increased platelet activity and thrombin generation seen directly after the operation in this study. However, all groups in the study have been treated according to the same routine, and we therefore believe that comparisons between the groups can be made.
Activation of coagulation and inflammation during CPB is a complex interplay involving both the intrinsic and, to a larger extent, the TF pathways. In experimental studies with simulated extracorporeal circulation, thrombin generation occurs before TF expression on monocytes, the only blood cell able to express TF, indicating a role of the intrinsic pathway. 38 In vivo, the surgical trauma with exposure of TF and involvement of inflammation makes the picture more complex.
Autotransfusion of shed blood after cardiac surgery is an often-used method to reduce the exposure of patients to homologous blood, 14,15 however debated. 17,22,39 Differences of threshold levels for transfusion between countries and clinics further complicates the possibility to evaluate and compare these studies. 40 In our study, in which no autotransfusion was performed, 30% of the patients needed homologous blood and received 2.2 units per patient, or a total of 0.67 units per patient. Furthermore, the quality of shed blood has been questioned, 39,41 with indications of defective hemostasis 19,22,42 and reports of systemic blood activation 21 and more febrile patients 39 after autotransfusion of this blood.
We found, in agreement with previous studies, that the SMB after CPB was characterized by activated cells, extraordinary activated coagulation, and inflammatory processes. The shed blood contained few cells but with a high expression of surface markers, indicating elevated activation. The low number of cells might be caused by adhesion, lysing, or dilution by means of exsudation.
In this study autotransfusion was not performed, and the duration of increased inflammatory and coagulant activity appeared to be shorter compared with that seen in previous studies in which shed blood was retransfused. 20,27,41,42 The high postsurgical levels of thrombin formation may be a consequence of the autotransfusion. Thrombin has been shown to induce several cell responses involved in inflammation, including, among others, cytokine release in monocytes and endothelial cells. 43 In contrast, the levels of IL-10, an anti-inflammatory cytokine and a potent inhibitor of monocyte activation and with anticoagulant effect, 44 did not differ between systemic blood and shed blood. Consequently, the imbalance between the cytokines in shed blood may reflect ongoing activity with subsequent effects on the coagulation and inflammatory systems.
During part 1 of the present study, we could conclude that the newly developed heparin surface is more biocompatible than an uncoated surface. The differences seen between parts 1 and 2 of the study are merely reflecting the importance of the systemic heparin level than the effect of the surface. The low dose of systemic heparin may not be sufficient to maintain the antithrombotic activity at the same level as a standard heparin dose during this kind of operation. The high dose, on the other hand, resulted in a direct cell-activating situation rather than a further anti-inflammatory and anticoagulatory effect.
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