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Original Articles

Tirofiban Administration Attenuates Platelet and Platelet-Neutrophil Conjugation but not Neutrophil Degranulation during In Vitro VAD Circulation

King, Bryan O.,*†; Whittow, Eluned S.H.,*†; Serna, Dan L.,‡; Jones, Blanding U.; Eng, Jamie S.M.,*; Chen, John C.*†

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Congestive heart failure (CHF), and its associated morbidity and mortality, continue to threaten millions of people in the United States. More than 400,000 new cases are diagnosed each year. 1 Treatment of end-stage heart failure is currently limited to heart transplantation and the use of mechanical ventricular assist devices (VAD). While the number of patients with end-stage heart failure who meet transplant criteria is increasing every year, the number of available donor organs remains substantially inadequate. 2

Cardiac support with VADs has improved survival in several patient groups with end-stage heart failure, 3 and VADs are now used for patients with postcardiotomy cardiogenic shock, as well as those awaiting transplantation. 4 Ventricular assist devices maintain systemic perfusion and serve as a bridge to heart transplantation. Unfortunately, the long-term use of VADs is accompanied by significant morbidity. 1 Clinical use of VADs continues to be associated with complications of bleeding, thromboembolism, infection, and end-organ failure. A number of immune cells and humoral mediators, such as platelets, neutrophils, and complement, have been implicated in VAD associated morbidity. 4 Platelet dysfunction may play a role in both bleeding and thromboembolic complications associated with prolonged use of the ventricular assist device. 1 Previous evidence suggests that VAD associated platelet dysfunction may be due to dysfunction of the platelet fibrinogen receptor. This receptor for fibrinogen is associated with the glycoprotein (GP) IIb/IIIa complex. 5 Neutrophils, which release elastase and form aggregates in the vasculature during extracorporeal circulation, also participate in the initiation and propagation of venous thrombosis. 6

In addition to thromboembolic complications, prolonged VAD use is also associated with an increased incidence of inflammation. The effects of blood-biomaterial contact on the complement system and the generation of inflammatory mediators have been well characterized in patients undergoing cardiopulmonary bypass. 7

The complement system is a collection of circulating proteins that together form a cascade of proteolytic activity. The complement system may become activated by one of two pathways, either the classical or alternative pathway, each of which converges into the common terminal pathway. Generation of C3a, the first activation fragment generated by the common terminal pathway, has been shown to occur early during in vitro VAD circulation. 1 Anaphylatoxins, C3a and C5a, along with the complement membrane attack complex (C5b-9), are potent leukocyte and platelet agonists 7 and may play a role in mediating the incidence of inflammation and thromboembolism. Platelets can regulate the classical pathway of complement activation via the secretion of C1 serine protease inhibitor (C1 INH), which is known to reside in platelet alpha granules. 8 C1 INH inhibits the proteases of the contact activation system and C1s and C1r, the initial proteases of the classical complement system. 1

The purpose of this study was to test the hypothesis that platelet and neutrophil activation are inter-related and linked to activation of the glycoprotein (GP) IIb/IIIa platelet receptor by evaluating the effects of GP IIb/IIIa receptor inhibition with Tirofiban on platelet and neutrophil activation during simulated VAD circulation. From the results of this study, the mechanisms of platelet and neutrophil activation may be further elucidated, and the role of complement in facilitating such activation during VAD circulation can be assessed.

Materials and Methods

Patient Selection

The protocol was approved by the Human Subject Institutional Review Board at the University of California, Irvine. Blood was obtained from volunteer human subjects. Subjects were asked to refrain from the use of any medication, including aspirin and other non steroidal anti-inflammatory medications for 2 weeks prior to donating blood. The selection process was random, and no restriction was made regarding the gender, racial/ethnic, age, or socioeconomic composition of the sample population.

In Vitro VAD Circuit

The simulated ventricular assist device circuit (surface area = 0.40 m2) consisted of a centrifugal pump (Medtronic Bio-Medicus pumphead, model # 9154R, Medtronic Blood Systems, Inc., Anaheim, CA, and Bio-Console, Medtronic Bio-Medicus, Inc., Eden Prairie, MN), polyvinyl chloride tubing (Medtronic Blood Systems, Inc.), and a polyvinylchloride venous reservoir bag (Baxter Health Care Corp., Fenwal Division, Maricao, Puerto Rico). Heparin sodium was obtained from Elkins-Sinn, Inc. (Cherry Hill, NJ).

Blood compartments of the assembled circuits were primed with 450 ml of plasmalyte (Baxter Health Care Corp., Santa Ana, CA). A total of 450 ml of blood was drawn from an antecubital vein into a venous reservoir bag containing heparin (2.5 units/ml). The venous reservoir bag was immediately attached to the VAD circuit and circulated into the VAD; each VAD circuit utilized one blood donor. A total of 10 blood donors were used during the course of this study. VAD circuits were used once and then discarded. Blood entered the VAD circuit from the venous reservoir bag by gravity (Figure 1).

Figure 1
Figure 1:
Schematic design of thein vitro VAD model.

Ten in vitro non pulsatile centrifugal VAD circuits were maintained for 3 days, each using 450 ml of fresh human whole blood. In order to assess the effects of platelet inhibition on platelet and neutrophil activity during VAD circulation, Tirofiban, a specific GP IIb/IIIa receptor inhibitor, was added to five of the VAD circuits. These circuits treated with Tirofiban formed the experimental group while the remaining five untreated circuits served as the control. Each experimental circuit received 50 μg of Tirofiban that was injected by a syringe into the venous reservoir bag immediately after the first sample was withdrawn for the baseline measurement. Because of the closed circuit design of this model, Tirofiban remained in the blood during the course of the 72 hour circulation period.

Blood was tested at regular time intervals using a blood gas analyzer (288 Blood Gas System, Ciba-Corning Diagnostics Corp., Medfield, MA), an ACT (Automated Coagulation Timer, Medtronic Hemotec, Inc., Englewood, CO), and a blood glucose meter (Lifescan, Inc., Milpitas, CA). Blood temperature was maintained at 37°C to simulate the in vivo circulating temperature of blood during clinical VAD use, using a heat exchange coil placed in a water incubator (Lab-Line Instruments, Inc., Melrose Park, IL. Activated clotting time, pH, PCO2, PO2, Ca2+, and glucose were maintained at physiologic levels. PO2 was maintained between 100 and 300 mm Hg by adjusting the rate at which oxygen was sterilely ventilated into the venous reservoir bag. Blood pH was maintained in a range of 7.2–7.6 using sodium bicarbonate, and PCO2 was maintained between 40 and 60 mm Hg by adjusting the ventilation rate of air into the venous reservoir bag. Blood flow was maintained at a cardiac index of 2.0 L/min per m2, because this rate provides adequate tissue perfusion during clinical use of VADs and cardiopulmonary bypass (CPB).

The first sample for analysis was drawn directly from the venous reservoir bag immediately prior to circulation in the VAD. Subsequent samples were drawn from a side port of the VAD circuit into 5 ml syringes at 10 min, 30 min, and 1, 2, 3, 4, 5, 6, 8, 12, 24, 48, and 72 hours after the start of blood circulation. Serum concentrations of platelet factor 4 (PF4), neutrophil elastase, and leukotriene C4 (LTC4) were measured using an enzyme linked immunosorbent assay (ELISA).

Enzyme Linked Immunosorbent Assays

Platelet factor 4 (PF4), neutrophil elastase, and leukotriene C4 (LTC4) assays each required 1 ml of blood to be drawn at each time interval. The first sample was drawn just prior to contact with the VAD surface. Another sample was withdrawn immediately after the blood entered the in vitro VAD circuit. Thereafter, samples were taken after 1, 2, 3, 4, 5, 6, 12, 24, 48, and 72 hours of VAD circulation. Immediately after withdrawal, blood samples were separated into 400 μl aliquots and combined with 40 μl of ethylenediaminetetraacetic acid (EDTA, 0.02 M). Samples were centrifuged for 15 min at 2000 g at 4°C. The supernatant (serum) was immediately collected and stored at −70°C. At the time of assay, frozen plasma samples were thawed to room temperature. Assays for the concentrations of LTC4 (Neogen Corp., Lansing, MI), PF4 (American Bioproducts Co., Parsippany, NJ), and Elastase (Merck, Inc., Whitehouse Station, NJ) were performed using enzyme linked immunosorbent assay kits.

PF4 Enzyme Linked Immunosorbent Assay

For PF4 assay, two dilutions were performed for each serum sample, one at 1:100 and the other at 1:1,000. For each standard, control, and sample dilution, 100 μl supernatant was mixed with 20 μl reconstituted PF4 reactant in a microfuge tube, and vortexed vigorously; the solutions were then all incubated for 30 min. An enzyme immunoassay plate reader read the microassay plate at a wavelength of 405 nm, and absorbances of the standards and controls were plotted with their known concentrations. The concentration of PF4 in the serum samples was then extrapolated from this plot.

Elastase Enzyme Linked Immunosorbent Assay

Dilutions (1:300 ) were performed for the samples by diluting 5 μl of sample with 1,495 μl of dilution medium. In addition, 1:100 dilution was performed for the baseline and 0 hour samples by diluting 20 μl of sample to 1,980 μl of dilution medium. A total of 500 μl of dilution medium (blank), calibrators, diluted control plasma, and diluted samples was transferred into tubes precoated with elastase α1-PI complex antibodies, and incubated for 1 hour. After incubation, the wells were washed three times with 1,000 μl of wash buffer to remove any unbound material. A total of 500 μl of a prepared antibody-enzyme solution marked with alkaline phosphatase was then added to each well and incubated for 30 min. Another three cycle wash was performed, but this time using 2,000 μl of wash buffer. A total of 500 μl of a freshly prepared substrate solution was then added to the wells and incubated in the dark for 30 min. Lastly, 100 μl of a stop solution was added to the tubes, and 200 μl of the solution in each tube was transferred into a microassay well. The microassay plate was read at 405 nm with an enzyme immunoassay plate reader. Known concentrations of the calibrators were plotted against their determined absorbances, and the concentrations of elastase in the plasma samples were then extrapolated from this plot.

LTC4 Enzyme Linked Immunosorbent Assay

Blood samples were initially centrifuged and extracted from serum using Sepharose column chromatography. The flow rate of the sample through the column was 1 ml per minute. Columns were washed with 1 ml of ethanol followed by 2 ml of distilled water, then eluted from the columns using 2 ml of methanol. The eluent was evaporated with nitrogen gas, leaving behind dry dissolved residue containing LTC4 that was then diluted and transferred into a microassay plate precoated with murine LTC4 complex antibodies along with 50 μl of diluted enzyme conjugate and incubated for 1 hour. After incubation, the wells were washed three times with 1,000 μl of wash buffer to remove any unbound material. A total of 150 μl of color substrate was then added to each well and the plate was incubated for 30 minutes, after which time the absorbance of the plate at 650 nm was spectrophotometrically measured. Absorbances of the sample wells were calculated using a standard concentration curve.


Leukotriene C4

In the group receiving Tirofiban, LTC4 levels increased to 120% of baseline after 3 hours of circulation and, thereafter, steadily declined (Figure 2). In the control group, LTC4 levels rapidly increased to 148% of baseline during the first hour of circulation. Thereafter, LTC4 levels remained elevated above baseline levels for the remainder of VAD circulation. The highest level of LTC4 in the control group was 158% over baseline, which occurred at 48 hours of VAD circulation. By 72 hours of circulation, LTC4 levels in the Tirofiban group declined to 23% of baseline versus 127% baseline in the control group.

Figure 2
Figure 2:
Percent baseline values of serum leukotriene C4 (LTC4) in VADs treated with Tirofibanversus untreated control VADs.


In both control and experimental groups, platelet release of PF4 appeared to occur continuously throughout in vitro VAD circulation without reaching a peak by 72 hours of circulation (Figure 3). PF4 levels in the control group steadily increased to 6.12 times the baseline value, from 652.90 IU/ml to 3,998.4 IU/ml, over 72 hours of in vitro VAD circulation. In VADs treated with Tirofiban, PF4 levels increased 3.19 times baseline value, from 1,003.66 IU/ml to 3,204.40 IU/ml, over 72 hours of circulation. These results indicate that Tirofiban reduced PF4 release from platelets to approximately half the amount of control VADs over the 72 hour course of in vitro recirculation. The differences in PF4 values between the treated and untreated groups were significant (p < 0.05) at 4, 5, 12, and 60 hours of circulation.

Figure 3
Figure 3:
Serum concentrations of platelet factor 4 (PF4) in VADs treated with Tirofibanversus untreated control VADs.


Elastase serum concentrations in the untreated VAD group increased from 126.5 μg/L at baseline to 925.02 μg/L at 24 hours and 2052.6 μg/L at 72 hours of in vitro VAD circulation (Figure 4). In VADs treated with Tirofiban, serum elastase increased from 191.73 μg/L at baseline to 1,356.68 μg/L at 24 hours and 1,836.29 μg/L at 72 hours of circulation. The differences observed between treated and untreated VADs were significant (p < 0.05) at 3, 4, 5, 6, and 12 hours. However, by 48 hours, both groups had risen to similar levels, after which they continued to increase in parallel up to 72 hours of circulation.

Figure 4
Figure 4:
Serum concentrations of human neutrophil elastase in VADs treated with Tirofibanversus untreated control VADs.


Ventricular assist devices are successfully utilized for postcardiotomy failure, postinfarction low-output syndromes, and as bridges to transplantation. 3,9,10 Depending on the pump design, VADs can deliver either pulsatile or non pulsatile flow. Although non pulsatile pumps are generally used for short-term support and pulsatile pumps for long-term support, current development of long-term non pulsatile VADs are under way and the superiority of either system remains undetermined. 11 Because non pulsatile circulatory support devices are widely used and available at most hospitals, we used the BioMedicus non pulsatile system as the basis for our in vitro VAD model.

Inherent to both types of flow systems is the effects of shear flow on platelets. 12,13 Several studies have demonstrated that shear flow induces platelets to express functional receptors that mediate platelet aggregation. 14,15 Previous studies by our group have shown that VAD induced aggregation may cause selective degradation of the platelet glycoprotein (GP) IIb/IIIa complex that likely causes insensitivity to aggregation agonists and leads to subsequent aggregation dysfunction. 8 Loss of platelet function during ventricular assist causes clinical complications such as prolonged bleeding and thrombogenesis which currently limits the prolonged use of ventricular assist devices. The glycoprotein (GP) IIb/IIIa receptor complex, a major GP of the platelet membrane, plays a significant role in platelet activation during ventricular assist. The receptor for fibrinogen is associated with the GP IIb/IIIa complex. Fibrinogen adsorbed onto the synthetic surfaces of the bypass circuit has been shown to activate platelets through a receptor, which is part of the GP IIb/IIIa receptor complex. 16 The platelet becomes adherent to the fibrinogen on the synthetic surface and platelets are then damaged by shearing forces of the flowing blood. They may become fragmented and lose their granule contents as well as their ability to aggregate. Thromboembolism after VAD use has been reported to occur in more than 30% of patients. 17–20

Despite this success of VADs, much remains to be learned about how these devices activate hematologic and inflammatory responses. Previous studies by our group have shown that addition of Tirofiban, a specific GP IIb/IIIa receptor inhibitor, attenuated platelet activation during in vitro ventricular assist and preserved the circulating platelet population. 5 Findings from this study indicate that Tirofiban also reduced the formation of neutrophil-platelet (Neut-Plt) conjugates in our in vitro VAD model. Although the physiologic consequences associated with these conjugates remain largely undetermined, recent studies have found the levels of circulating conjugates doubled before thromboembolic neurologic events in VAD patients, implying a role for these conjugates in precipitation of stroke. 7 The production of LTC4 suggests Neut-Plt conjugation occurs during in vitro VAD circulation. Generation of LTC4, a potent inflammatory mediator, occurs from the collaboration between neutrophils and platelets resulting in the transcellular biosynthesis of LTC4. 21–23 The path to LTC4 production begins in the activated neutrophil with the liberation of arachidonic acid from phospholipid precursors, and the generation of LTA4 by 5-lipoxygenase within the neutrophil. When platelets are present, this LTA4 passes to platelets where it is converted to LTC4 by the platelet glutathione-S-transferase. If platelets are not present, the LTA4 within neutrophils will usually be hydrolyzed nonenzymatically or converted to LTB4. Neither platelets nor neutrophils alone possess both enzymes required to produce LTC4. This process is enhanced by the close proximity of neutrophils and platelets and by the combined activation of both neutrophils and platelets, both of which may release phospholipid precursors that may participate in the production of LTC4. This cooperative process has been demonstrated in whole blood. 24

Unfortunately, Neut-Plt conjugation may result in a variety of harmful sequelae. Neut-Plt conjugates may embolize to distant organs and cause ischemic events and end-organ failure. 25 Furthermore, LTC4, along with LTD4 and LTE4, are the components that make up the slow reacting substance of anaphylaxis (SRS-A). 24 LTC4 can cause platelet aggregation, bronchoconstriction, vasoconstriction, increased vascular permeability, early pulmonary changes characteristic of adult respiratory distress syndrome, and hemodynamic changes characteristic of anaphylaxis and sepsis. 26 Decreased LTC4 with extended VAD circulation suggests decreased Neut-Plt conjugation. With the addition of Tirofiban, there is decreased Neut-Plt conjugation at 72 hours of circulation in our in vitro VAD model. Our findings support a mechanism by which platelets play a regulatory role in mediating conjugate formation during VAD use.

In addition to thromboembolic complications, prolonged VAD use is also associated with an inflammatory response. 7 Our findings demonstrate that neutrophils are activated and release their granule contents during in vitro VAD circulation. Neutrophil activation results in the release of proinflammatory enzymes, including myeloperoxidase, lactoferrin, elastase, and other lysosomal hydrolases from secretory azurophilic granules. These granule proteins aid in the formation of oxygen free radicals, which damage a variety of cellular components. During cardiopulmonary bypass, oxygen free radical damage has been associated with ischemia, reperfusion mediated myocardial injury, reperfusion arrhythmia, and postoperative cardiac dysfunction. 27 Furthermore, neutrophils participate in the initiation and propagation of venous thrombosis. Human elastase is proteolytic and can degrade extracellular matrix molecules such as fibronectin, proteoglycans, and type IV collagen. Elastase, when present in abnormally high concentrations, initiates tissue destructive reactions, the effects of which contribute to pulmonary emphysema and certain forms of liver disease. Proteolytic enzymes such as elastase must normally be closely regulated and quickly inactivated before overexposure to such enzymes causes tissue damage. The observed increase in elastase levels during VAD use would explain some of the inflammatory and tissue destructive complications during ventricular assist. In patients utilizing left ventricular assist device support, it has not been elucidated to what extent mechanical circulatory support, as opposed to the underlying condition of end-stage heart failure, is responsible for increasing serum levels of proinflammatory mediators.

The complex interaction between platelets, complement, and inflammatory mediators such as neutrophils make it difficult to determine the chronologic sequence by which these factors are activated. Previous studies by our group demonstrated that VAD induced activation of platelets stimulated the release of C1-Inhibitor (C1-INH), a serine protease inhibitor stored within platelet alpha granules that regulates the classical pathway of complement. 1 Compared with control VADs, Tirofiban treated VADs demonstrated significantly lower serum levels of C1-INH during the first 6 hours, after which time C1-INH increased to levels > 150% of control for the remainder of the experiment. In this study, neutrophil activation in the Tirofiban group was initially higher than the control group, as demonstrated by higher levels of elastase. The initially higher levels of neutrophil activity in Tirofiban treated VADs is associated with reduced complement regulation caused by decreases in serum C1-INH. After 6 hours, C1-INH levels in Tirofiban treated VADs increased above control levels, correlating temporally with the observed decreased rate of elastase release in the experimental group. No significant difference was observed in elastase levels after six hours of circulation. With the exception of initial circulation, the finding that platelet inhibition does not affect neutrophil activation during the majority of VAD circulation indicates that platelets may not play the predominant regulatory role in mediating inflammation. Rather, it is more likely that complement, specifically C5a, precedes platelet activation and is responsible for neutrophil activation. C5a, a potent anaphylactic neutrophil agonist, is produced from the terminal complement cascade (TCC) and has been shown to cause neutrophil elastase release. Recent studies by Rinder et al.28,29 using simulated extracorporeal circulation (SECC) appear to support the role of C5a in stimulating neutrophil activation. Their studies demonstrated that among the biologic components produced by the TCC (C3a, C5a, and C5b-9), only C5a directly contributed to neutrophil activation during SECC. By inhibiting C5a and C5b-9 (also known as the membrane attack complex or MAC) during separate experiments, they found that inhibiting the formation of MAC did not affect neutrophil activation, while inhibition of C5a significantly reduced neutrophil CD11b upregulation.

The early activation of complement is not only likely to precede platelet activation, but may, in fact, play a role in platelet activation as well as formation of platelet-neutrophil conjugates. C5a stimulates the release of other substances stored in the azurophilic granules such as cathepsin G, a serine protease that induces platelet calcium mobilization, aggregation, and serotonin release. 30 Thus, a pathway producing thrombosis exists whereby neutrophils can directly activate platelets, thereby inducing aggregation and thrombus formation. Conversely, platelets also have the ability to directly activate neutrophils. Platelet factor 4 (PF4), which resides in platelet alpha granules, is known to induce neutrophil activation and degranulation. Platelets, by degranulation of PF4, may, therefore, play a role in causing initial neutrophil activation and a significant role in mediating platelet-neutrophil conjugation during clinical ventricular assisted circulation.

In conclusion, it is likely that the onset of blood interaction with the artificial surfaces of the VAD is characterized by early complement activation. Subsequent formation of C5a and the MAC causes platelet activation and conjugate production, while C5a alone stimulates neutrophil activation. Complement activation appears to occur early and play a significant role in mediating both the inflammatory and thromboembolic events observed in VAD patients. However, several feedback mechanisms exist that appear to allow amplification of the initial complement stimulus. Such feedback systems include the direct regulation of conjugate formation by platelets and the ability of neutrophils to activate platelets via the release of granule components. Therefore, future studies should be directed toward pharmacologic attenuation of C5a and MAC in blood circulating through ventricular assist devices in order to further elucidate the biochemical mechanisms causing inflammatory and thromboembolic reactions that currently plague the long-term use of ventricular assist devices.


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