Platelet adhesion to surface-bound protein on the subendothelium is the initial condition
of blood coagulation. Following adhesion, the platelets become activated and in turn generate thrombin and thromboxane A2
on their extracellular membranes and release ADP from intracellular stores to promote further platelet activation and aggregation. The release of these platelet agonists into the blood flowing over the adhered platelets is the boundary condition
that leads to platelet aggregation. The flux of these three agonists into flowing blood serves an important role in thrombosis and hemostasis. However, the combined function of such soluble agonists during clot formation under flow is unknown, since no method allows controlled fluxes of agonists into flowing blood. The relationship between solute flux and stable clot formation is a complex reaction–transport problem with coupled particle deposition. One example of altered solute flux is hemophilia, where deficiencies in clotting factors attenuates thrombin generation,26
which leads to unstable clots.27
It is less clear whether these clots are unstable because of diminished fibrin formation or diminished platelet activation, or both. The device described in this paper provides a method for manipulating solute flux that could be useful for studying hemophilia and other bleeding disorders.
The size and growth of a thrombus is limited by the interplay between activated platelets and intact endothelial cells. Endothelial cells secrete soluble molecules (nitric oxide, prostacyclin, ectonecleotidases) that control platelet reactivity in a process known as thromboregulation.28
The device presented in this paper could be extended to study thromboregulation with additional bottom channels for introduction of these platelet antagonists.
Incorporation of membranes into microfluidic devices is becoming increasingly common for on-chip filtration and separations, as well as a component for sensors, reactors, and cell culture.29
In this paper we incorporated commercially available membranes as fluidic resistor into a microfluidic device by modifying a technique first reported by Ismagilov and colleagues.30
This technique has been employed for screening biochemical interaction,30
generating chemical gradients,31
and monitoring communication between different cell types.32
A notable modification we have made was the use of vacuum assisted bonding.33
This reversible bonding technique allowed for post hoc
morphological analysis of platelet aggregates by electron and spinning disk confocal microscopy.
Track-etched polycarbonate membranes demonstrate both internal and external fouling under different operating conditions.24,25,34
Internal fouling describes the adsorption and deposition of molecules and particles onto the surface of the pore, and consequently reducing the pore size. External fouling describes the accumulation of larger particles and cells that cannot enter pores. We have chosen a membrane whose pores are much larger than any of the solutes we are interested in introducing into blood, but also small enough to exclude blood cells. However, we still observed fouling of the pores that led to significant flux decline over 25 min. This flux decline was most likely the result of internal fouling based on moderate transmembrane pressures and short length of the experiments.25
Other choices of membranes include cellulose acetate, polyvinylidene fluoride, or polysulfone. However, these other membranes have a minimum thickness on the order of 100’s microns, whereas the polycarbonate membranes are 5–15 μm thick. The membrane thickness is an important parameter in the multilayer device described in the paper. We attempted to use a cellulose acetate membrane (thickness = 110 μm) and found that the loss of fluid into the vacuum chamber was prohibitive.
Solute transport was accurately predicted with an analytical model when convection was the dominant transport mechanism within the pores (Pe > 10). Although the model did not account for protein fouling, it was a good first-order etstimate of flux that only required two parameters, the membrane permeability and transmembrane pressure. The device was operated in two configurations that depended on the wall shear rate in the top channel. At higher wall shear rates (1000–2000 s−1), there was a large enough transmembrane pressure generated by the top channel to achieve predominantly convective transport in the pores. At these high shear rates, fluid was withdrawn (negative pressure) from both channels. At lower wall shear rates (250–500 s−1), the transmembrane pressure generated by the top channel was insufficient to ensure convective transport. Therefore, at these low shear rates, the bottom channel was infused (positive pressure) to compensate for small pressure drop in the top channel. In some applications, such as gradient generation, diffusion-mediated transport may be preferable. This mode of operation is achievable by withdrawing fluid from both channels at the same flow rate, provided the channels have the same length and cross-sectional area.
As a demonstration of the device, we examined the dependence of platelet aggregation on ADP flux. We observed that platelet activation, aggregation size, and aggregation height depended on the magnitude of ADP flux. At the lowest ADP flux considered, there was sparse adhesion of platelets and little evidence of aggregation and platelet–platelet tethering. At the higher fluxes, we observed aggregates of hundreds of activated platelets. The size and height of aggregates increased with increasing ADP flux. These observations agree with previous reports that platelet adhesion efficiency to fibrinogen coated surfaces increases with ADP induced activation.23
In addition, computational studies predict that a thicker agonist boundary layer yields greater platelet aggregation.35–37
In these computational studies, boundary layer thickness and thus platelet aggregation was modulated by changing the wall shear rate. However, increasing the agonist flux has a similar effect to decreasing the wall shear rate.