Name: microfluidics in assessing platelet function
Uploaded: 2024-11-08T14:10:00.000+00:00
Duration: 6 min 47 s
Description: Microfluidics incorporate physiologically relevant substrates and flows that mimic the vasculature and are, therefore, a valuable tool for studying ...

The authors acknowledge and thank all blood donors who participated, as well as the Trauma and Transfusion Medicine Research Lab phlebotomists and the UPMC Montefiore Clinical and Translational Research Center for assistance in collections. SMS is supported by K25HL161401. MDN is supported by 1R01HL166944-01A1.

All research was performed in compliance with institutional guidelines. Approval from the University of Pittsburgh Human Research Protection Office was obtained and informed consent from healthy human volunteers was obtained.
1. Microfluidic device preparation
<ol>
	<li>To fabricate the PDMS portion of the device, prepare a master mold using brass via computer numerical control (CNC) micromachining. 
	NOTE: Depending on channel dimensions, photolithography techniques may be used to create a master mold. The device used in this protocol includes eight parallel micromachined channels approximately 480 &#181;m wide, 140 &#181;m tall at the device inlet and outlet, and 40 &#181;m tall at the device stenosis, with a ramp length to/from the stenotic region of approximately 0.3 mm. Channel lengths are approximately 6 mm.</li>
	<li>With a master mold obtained, pour the Silicone Elastomer Base (obtained from the elastomer kit) into a weighing dish. Add the Silicone Curing Agent (obtained from the elastomer kit), which facilitates the cross-linking of the silicone polymer chains to transform the liquid PDMS into a durable and flexible solid, at a ratio of 10:1 (base agent) and stir the mixture well.</li>
	<li>Place the mold into a Petri dish and pour the uncured PDMS onto the mold. Place the Petri dish inside a vacuum desiccator for 30 min to remove bubbles.</li>
	<li>Finish curing the PDMS in the master mold by placing it in an oven set to 70 &#176;C for 90 min.</li>
	<li>After complete PDMS curing, cut out the microfluidic cast using a razor blade or scalpel. Punch holes at the edges of the channels (1.5 mm in diameter on both sides).</li>
	<li>Using lab tape, clean the surface of a glass slide and the etched side of the microfluidic cast. Use compressed air as needed to remove remaining debris.</li>
	<li>Place the glass slide and the microfluidic cast with etched side upward into a Plasma Cleaner. Start the vacuum pump, seal the chamber, and turn on the plasma cleaner to the high setting. Leave the slide and PDMS in the plasma cleaner for 30 s, then turn the plasma cleaner off and remove vacuum.</li>
	<li>Bind the plasma-cleaned cast and glass slide together by gently pressing together the sides that were face up in the plasma cleaner. Then, place the microfluidic device in an oven/hotplate at 70 &#176;C for 10 min. 
	NOTE: Do not apply too much pressure when bonding the cast and glass slide together as that could lead to a loss in channel morphology.</li>
	<li>Rinse each chamber with 10-30 &#181;L of 70% ethanol to sterilize and allow the microfluidic device to dry on a 70 &#176;C hot plate. 
	NOTE: Devices should be made at least 24 h in advance but may be made weeks to months in advance. Devices should be stored in an airtight container or covered Petri dish at room temperature.</li>
	<li>The day before experimentation with the microfluidic device, re-rinse each chamber with 10-30 &#181;L of 70% ethanol to sterilize and allow the microfluidic device to dry on a 70 &#176;C hotplate.</li>
	<li>Coat the chamber with Type 1 equine fibrillar collagen reagent (1 mg/mL), diluted in 0.9% NaCl in a 1:5 volumetric ratio through a designated outlet to designated inlet. Make sure directionality is conserved within an experiment. Store the device in a warm, humid closed container to prevent evaporation of the coated collagen within the channel.</li>
	<li>After 1 h, rinse with phosphate-buffered saline (PBS) to flush out the collagen solution. Flush in the opposite direction of the coating. When not in use, store the device again in a warm, humid, closed container.</li>
</ol>
2. Blood sample preparation
<ol>
	<li>Obtain a citrated whole blood sample via venipuncture. Incubate citrated whole blood with FC receptor blocking solution (1:600). 
	NOTE: Blood samples are stored at room temperature prior to and during experimentation.</li>
	<li>Incubate citrated whole blood with fluorescence-conjugated (using a fluorophore of choice) CD41 antibody (1:600). Stain for 30 min on a nutating rocker.</li>
	<li>As a positive control for platelet inhibition, add Ticagrelor, a P2Y12 receptor antagonist reconstituted in a solution of 30% w/v 2-hydroxypropyl-&#946;-cyclodextrin (HP-&#946;-CD) in PBS. 
	NOTE: Ticagrelor stocks up to 6.4 mM should be prepared and further dilution of Ticagrelor stocks in HP-&#946;-CD solution (30% HP-&#946;-CD dissolved in PBS) can be done prior to experimentation (1,000x stock concentrations are recommended).</li>
	<li>Incubate the citrated sample with Ticagrelor (final concentration up to 6.4 &#181;M) for 30 min to observe platelet inhibition.</li>
	<li>If mixing the blood product with the citrated sample, stain the blood product with FC receptor blocking solution (1:600) and fluorescence-conjugated (using a separate and distinct fluorophore to the citrated sample) CD41 antibody (1:600).
	<ol>
		<li>Mix a volumetric equivalent of the transfused product units with the citrated sample. For example, to simulate 2 units of platelet products transfused into a hemorrhaging person (approximately 250 mL per product into a total blood volume of 5,000 mL), mix 100 &#181;L of blood product into 1,000 &#181;L of citrated blood sample.</li>
	</ol>
	</li>
	<li>Immediately before experimentation, filter the blood sample through a 40 &#181;m filter into a sterile 1.5 mL microcentrifuge tube.</li>
</ol>
3. Platelet function testing under flow (Method 1)
<ol>
	<li>Turn on the microscope and associated software.</li>
	<li>Set a withdrawal syringe pump level with the stage of the microscope. Adjust the settings on the syringe pump.
	<ol>
		<li>Calculate a volumetric flow rate (Q) for a desired average wall shear rate (&#947;) of 3,500 s-1 at the stenotic region of the channel using equations (1) and (2)21.&#160;&#160; 
		<img alt="Equation 1" src="/files/ftp_upload/67214/67214eq01.jpg" style="margin: auto;" />&#160; &#160; &#160;(1) 
		<img alt="Equation 2" src="/files/ftp_upload/67214/67214eq02v2.jpg" style="margin: auto;" />&#160; &#160; &#160;(2) 
		Where A is the cross-sectional area of the channel, P is the wetted perimeter, &#955; is the shape factor, b is the short side of the rectangle (height), and a is the long side of the rectangle (width). 
		NOTE: The value of 3,500 s-1 is chosen due to VWF critical shear and is in the arterial regime7,22,23.</li>
	</ol>
	</li>
	<li>Place the microfluidic device on the microscope stage. Tape the edges of the microfluidic device to the stage to avoid movement. Ensure that the outlet is facing the back of the microscope.</li>
	<li>Connect one end of 1/16&#8221; ID tubing (approximately 30 cm long) to an elbow connector and the other end to a 10 mL syringe with 1/16&#8221; ID connector.</li>
	<li>Fill the syringe with sterile PBS and connect it to the syringe pump.</li>
	<li>Place the elbow connector into the device outlet.</li>
	<li>Prepare inlet lines approximately 10 cm long with the elbow connector on one end and the angled cut on the other end.</li>
	<li>Connect the inlet elbow connector into the device inlet.</li>
	<li>Position the inlet line into a waste microcentrifuge tube on an angled holder.</li>
	<li>Use the 10x objective for channel dimensions approximately 500 &#181;m in width and focus on the microfluidic device channel edges.</li>
	<li>Prime the lines with PBS and clear the channel of any PDMS/debris by advancing the syringe pump manually. Be sure to check near the channel inlet and outlet.</li>
	<li>Open the saved image capture settings or create an image capture procedure for time series images captured every 1&#8211;2 s with a brightfield channel and fluorescent channels corresponding to the fluorescent CD41 antibodies used in the blood sample.</li>
	<li>Obtain the filtered citrated blood sample and mix by pipetting up and down just prior to the experiment. Place the sample on the angled microcentrifuge holder.</li>
	<li>Position the inlet line into the sample.</li>
	<li>Start recording the image capture.</li>
	<li>Slowly, withdraw the syringe to fill the dead volume in the tubing. Once blood reaches the channel, immediately hit play on the syringe pump to resume flow at the desired shear rate.</li>
	<li>Adjust the focus if needed.</li>
	<li>Run the experiment until platelets have completely occluded the stenotic region of the microfluidic device or until an experimental endpoint (i.e., 10 min).</li>
	<li>Ensure the inlet tubing is submerged in the blood sample for the duration of the experiment.</li>
	<li>Once the experiment is complete, stop the image capture and stop the syringe pump. Save the image capture.</li>
	<li>Remove the inlet elbow connector and empty the contents of the tubing into a waste conical. Empty the contents of syringe and outlet lines into waste conical as well.</li>
	<li>Replace the inlet and outlet connectors and tubing as necessary for subsequent samples. &#160;</li>
</ol>
4. Platelet function testing under flow with low-volume samples (below 1 mL) (Method 2)
<ol>
	<li>Repeat steps 1.1 through 1.4&#160;as above.</li>
	<li>Punch a 1.5 mm in diameter outlet and 3 mm in diameter inlet at the edges of the channels.</li>
	<li>Repeat steps 1.6 through 3.6 as above.</li>
	<li>Use the 10x objective for channel dimensions approximately 500 &#181;m in width and focus on the microfluidic device channel edges.</li>
	<li>Clear the channel of any PDMS/debris by advancing the syringe pump manually and wicking away access fluid with a laboratory wipe. Remove any debris with lab tape on the top of the microfluidic device.</li>
	<li>Manually advance the pump to fill the 3 mm inlet reservoir with PBS.</li>
	<li>Open the saved image capture settings or create an image capture procedure for time series images captured every 1&#8211;2 s with a brightfield channel and fluorescent channels corresponding to the fluorescent CD41 antibodies used in the blood sample.</li>
	<li>Start withdrawal on the syringe pump (manually or with set speed) and once PBS is advancing into the channel and the fill line in the reservoir is going down, pause the withdrawal on the pump.</li>
	<li>Remove excess PBS from the reservoir with a pipette.</li>
	<li>Start the image capture.</li>
	<li>Pipette the blood sample into the reservoir (approximately 40 &#956;L) and immediately start the withdrawal syringe. Ensure flow starts. 
	NOTE: If the syringe pump motor for withdrawal was not primed in step 4.8, flow will not start immediately, and the experiment should be repeated.</li>
	<li>Refill the blood reservoir for the duration of the experiment, ensuring no air pockets enter the channel.&#160;</li>
	<li>Run the experiment until the platelets have completely occluded the stenotic region of the microfluidic device or until an experimental endpoint (i.e., 10 min).</li>
	<li>Once the experiment is complete, stop image capture and stop the syringe pump. Save the image capture.</li>
	<li>Remove excess blood in the reservoir by pipetting. Empty the contents of the withdrawal tubing into a waste conical tube.</li>
	<li>Replace the connectors and tubing as necessary for subsequent samples.&#160;</li>
</ol>
5. Decontamination
<ol>
	<li>Clear the inlet and outlet lines of blood by flushing into a 10% bleach solution.</li>
	<li>If all the channels on the microfluidic device have been used, then discard the device in the Sharps biohazardous waste container. 
	NOTE: All biological waste should be disposed of in biohazard waste appropriately.</li>
</ol>
6. Image analysis
<ol>
	<li>Export the images from kinetic experiments using software by clicking File | Export/Import | Export.</li>
	<li>Select the following parameters: filetype: Tagged Image File Format (TIFF); compression: LZW; check Original Data and Shift Pixel; uncheck Apply display curve and channel color; check Define Subset (Region, Rectangle region, and select the channel area with a conserve width and height measurement between conditions). Export to the desired folder and check Create folder.</li>
	<li>Note the below experimental values on the software: start frame when blood enters the channel; frame rate (Info, Time Series).</li>
	<li>Measure the normalized mean fluorescence intensity of each frame in an experiment utilizing Matlab code provided as Supplemental File 1, changing the following inputs for each experiment: start frame/end frame based on experiment length (ensure the experiment length is conserved between conditions); experiment name/folder name; cropping X, Y, H, and W parameters. Run the code once for the recipient platelet fluorescent channel and once for the product platelet fluorescent channel. Report normalized MFI (column 2) and normalized AUC values (normalized to start frame). Subtract the experiment length value from the normalized AUC value.</li>
</ol>

Platform for Assessing Platelet Function in Trauma and Transfusion

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The authors have no conflicts of interest to declare.

The above protocol has some critical steps to ensure the reliability and reproducibility of experiments. First, fluorescent antibodies should be carefully considered. The antibodies used to detect platelets in the sample should not block the function of the glycoprotein Ib (GPIb) platelet receptor. Lot matching, whenever possible between experiments, is also critical to ensure the reproducibility of the fluorescent signal. Another critical step in this protocol is using sterile consumables and solutions and filtered samp...

Trauma is a leading global cause of death and disability. Severe injury is frequently complicated by a unique, endogenous disturbance of hemostasis and thrombosis, termed trauma-induced coagulopathy (TIC)1. Platelets play a critical role in TIC, and they have been described as having both adaptive and maladaptive functions2. The mechanisms of platelet dysfunction after injury remain unclear, and there is a critical need to better understand the cellular response to guide the development of improved resuscitation and therapy. An additional vexing problem regarding platelet function after injury is the uncertainty of the reliability of present readouts of platelet function in the trauma patient.
Multiple studies have shown that even mildly injured patients, with no known clinical bleeding phenotype, have abnormal platelet function using conventional platelet function testing such as aggregometry3,4. However, limitations in aggregometry to assess platelet function in an injury setting include a lack of physiologically relevant injury surface, a reductionist approach to agonist stimulation, sample dilution with whole blood impedance aggregometry, plasma separation with optical light transmission aggregometry, and stagnant sample assessment. Additionally, whether this sensitivity of platelet function represents true cellular dysfunction or a measurement artifact, such as increased baseline electrical impedance, in the setting of injury remains unclear2. Thus, studying relevant platelet functions in the context of trauma is crucial to understanding TIC, and there is substantial room for innovation and improvement in this area.
Platforms traditionally used to study platelet function do not include fluid dynamics and flow, which may be critical in understanding platelet dysfunction pertaining to trauma and trauma-induced coagulopathy5. Mechanisms of hemostasis that are dependent on flow include von Willebrand factor (VWF) elongation at high shear, above a critical shear rate, and platelet capture via glycoprotein 1b6,7,8, which are not captured using stagnant platelet function assays. Additionally, platelets preferentially bind VWF or fibrinogen depending on the flow regime and elicit differential roles in arterial versus venous thrombosis9,10. Arterial thrombi are mainly comprised of platelets while venous thrombi are mainly comprised of red blood cells, based, in part, on flow regimes11. Assays that incorporate flow regimes can aid in elucidating dysfunctions pertaining to the spectrum of TIC phenotypes, from hypocoagulability and bleeding phenotypes to hypercoagulability and thrombotic phenotypes. Finally, blood volume sampling constraints with trauma patient populations may make traditional platelet function testing challenging. While assays such as flow cytometry can and should be utilized in these circumstances, results often depict a physical characterization of a sample and not a hemostatic functional assessment.
While mechanisms of platelet dysfunction may not be completely understood in trauma, modeling platelet dysfunction in vitro, with P2Y12 antagonists for example, can also help guide the study of therapeutic interventions. Hemostatic resuscitation is critically important in trauma patients where blood products are transfused in a balanced approach to address shock, coagulopathy, and endothelial injury with either whole blood or blood components (red blood cells, plasma, and platelet concentrates) in a 1:1:1 unit ratio12,13,14. In trauma patients, early use of blood products is associated with improved survival15,16. To extend shelf life, cold-stored platelet products have been increasingly studied. Examination of cold-stored platelets shows increased hemostatic activity, as well as safety when transfused following injury17,18.
The evolution of cold-stored platelet resuscitation emphasizes the need for additional testing to understand the most efficacious platelet product available for trauma. However, traditional platelet function assays are often over- or under-potentiated to detect dysfunction, occurring both in the trauma patient receiving therapeutic platelet transfusion as well as in the transfused product itself seen in platelet storage lesions. Determining the origin of dysfunction can be challenging, given the limitations in current platelet function assays, including the static nature of most of these tests. Therefore, when studying hemostatic resuscitation in vitro, the platform and detection methods for both recipient and product platelet populations are of critical importance in determining optimal therapeutic interventions.
Microfluidic testing offers flow profiles and biofidelic surfaces to create a physiologically relevant assay on which to study platelets. Microfluidic devices can be customized to study particular pathophysiology or injury types, such as vessel puncture19 or endothelial damage20. These devices are generally comprised of polydimethylsiloxane (PDMS) bonded to a glass microscope slide with surface modifications, such as collagen, to mimic sub-endothelium and tissue injury. Utilizing these types of flow-based devices can aid in guiding trauma-related platelet dysfunction research and aid in examining optimal transfusion medicine approaches to ameliorate platelet dysfunction. These strategies may help to clarify the existing confusion about the relevance of static platelet assays such as aggregometry in the injured patient.

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microfluidics in assessing platelet function

Microfluidics incorporate physiologically relevant substrates and flows that mimic the vasculature and are, therefore, a valuable tool for studying aspects of thrombosis and hemostasis. At high-shear environments simulating arterial flow, a microfluidic assay facilitates the study of platelet function, as platelet-rich thrombi form in a localized stenotic region of a flow channel. Utilizing devices that allow for small sample volume can additionally aid in evaluating platelet function under flow from volume-limited patient samples or animal models. Studying trauma patient samples or samples following platelet product transfusion may aid in directing therapeutic strategies for patient populations in which platelet function is critical. Effects of platelet inhibition via pharmacological agents can also be studied in this model. The objective of this protocol is to establish a microfluidic platform that incorporates physiologic flow, biological surfaces, and relevant hemostatic mechanisms to assess platelet function with implications for the study of trauma induced coagulopathy and transfusion medicine.

Microfluidic experiments following the use of this method should show platelet-rich thrombi formation in the region of stenosis of the flow channel (Figure 1). Figure 1A illustrates representative results where functional platelets formed a thrombus in the stenotic region of the channel to block blood flow through the channel. Mean fluorescence intensity (MFI) curves of kinetic images taken for the duration of the experiment illustrate a lag, growth, and plateau...

Microfluidics in Assessing Platelet Function

Platelet function under flow can be assessed and simulated hemostatic resuscitation can be modeled using a microfluidic device, which has applications in trauma and transfusion medicine.

Microfluidics incorporate physiologically relevant substrates and flows that mimic the vasculature and are, therefore, a valuable tool for studying ...

microfluidics-in-assessing-platelet-function

Research

JoVE Journal

Medicine

738 Views.  University of Pittsburgh. Platelet function under flow can be assessed and simulated hemostatic resuscitation can be modeled using a microfluidic device, which has applications in trauma and transfusion medicine. 

Video: Microfluidics in Assessing Platelet Function

Assessing Endothelial Vasodilator Function with the Endo-PAT 2000

The endothelium is a delicate monolayer of cells that lines all blood vessels, and which comprises the systemic and lymphatic capillaries. By virtue of the panoply of paracrine factors that it secretes, the endothelium regulates the contractile and proliferative state of the underlying vascular smooth muscle, as well as the interaction of the vessel wall with circulating blood elements. Because of its central role in mediating vessel tone and growth, its position as gateway to circulating immune cells, and its local regulation of hemostasis and coagulation, the the properly functioning endothelium is the key to cardiovascular health. Conversely, the earliest disorder in most vascular diseases is endothelial dysfunction.
In the arterial circulation, the healthy endothelium generally exerts a vasodilator influence on the vascular smooth muscle. There are a number of methods to assess endothelial vasodilator function. The Endo-PAT 2000 is a new device that is used to assess endothelial vasodilator function in a rapid and non-invasive fashion. Unlike the commonly used technique of duplex ultra-sonography to assess flow-mediated vasodilation, it is totally non-operator-dependent, and the equipment is an order of magnitude less expensive. The device records endothelium-mediated changes in the digital pulse waveform known as the PAT ( peripheral Arterial Tone) signal, measured with a pair of novel modified plethysmographic probes situated on the finger index of each hand. Endothelium-mediated changes in the PAT signal are elicited by creating a downstream hyperemic response. Hyperemia is induced by occluding blood flow through the brachial artery for 5 minutes using an inflatable cuff on one hand. The response to reactive hyperemia is calculated automatically by the system. A PAT ratio is created using the post and pre occlusion values. These values are normalized to measurements from the contra-lateral arm, which serves as control for non-endothelial dependent systemic effects. Most notably, this normalization controls for fluctuations in sympathetic nerve outflow that may induce changes in peripheral arterial tone that are superimposed on the hyperemic response.
In this video we demonstrate how to use the Endo-PAT 2000 to perform a clinically relevant assessment of endothelial vasodilator function.

A noninvasive procedure to assess endothelial function is demonstrated using the Endo-PAT 2000.

The endothelium is a delicate monolayer of cells that lines all blood vessels, and which comprises the systemic and lymphatic capillaries. By virtue of ...

In Vivo Canine Muscle Function Assay

We describe a minimally-invasive and reproducible method to measure canine pelvic limb muscle strength and muscle response to repeated eccentric 
contractions. The pelvic limb of an anesthetized dog is immobilized in a stereotactic frame to align the tibia at a right angle to the femur. Adhesive wrap affixes the 
paw to a pedal mounted on the shaft of a servomotor to measure torque. Percutaneous nerve stimulation activates pelvic limb muscles of the paw to either push (extend) or 
pull (flex) against the pedal to generate isometric torque. Percutaneous tibial nerve stimulation activates tibiotarsal extensor muscles. Repeated eccentric (lengthening) 
contractions are induced in the tibiotarsal flexor muscles by percutaneous peroneal nerve stimulation. The eccentric protocol consists of an initial isometric contraction 
followed by a forced stretch imposed by the servomotor. The rotation effectively lengthens the muscle while it contracts, e.g., an eccentric contraction. During 
stimulation flexor muscles are subjected to an 800 msec isometric and 200 msec eccentric contraction. This procedure is repeated every 5 sec. To avoid fatigue, 4 min rest 
follows every 10 contractions with a total of 30 contractions performed.

In Vivo Canine Muscle Function Assay

We describe a minimally-invasive and painless method to measure canine hindlimb muscle strength and muscle response to repeated eccentric contractions.

We describe a minimally-invasive and reproducible method to measure canine pelvic limb muscle strength and muscle response to repeated eccentric 
...

Endothelialized Microfluidics for Studying Microvascular Interactions in Hematologic Diseases

Advances in microfabrication techniques have enabled the production of inexpensive and reproducible microfluidic systems for conducting biological and biochemical experiments at the micro- and nanoscales 1,2. In addition, microfluidics have also been specifically used to quantitatively analyze hematologic and microvascular processes, because of their ability to easily control the dynamic fluidic environment and biological conditions3-6. As such, researchers have more recently used microfluidic systems to study blood cell deformability, blood cell aggregation, microvascular blood flow, and blood cell-endothelial cell interactions6-13.However, these microfluidic systems either did not include cultured endothelial cells or were larger than the sizescale relevant to microvascular pathologic processes. A microfluidic platform with cultured endothelial cells that accurately recapitulates the cellular, physical, and hemodynamic environment of the microcirculation is needed to further our understanding of the underlying biophysical pathophysiology of hematologic diseases that involve the microvasculature.
Here, we report a method to create an "endothelialized" in vitro model of the microvasculature, using a simple, single mask microfabrication process in conjunction with standard endothelial cell culture techniques, to study pathologic biophysical microvascular interactions that occur in hematologic disease. This "microvasculature-on-a-chip" provides the researcher with a robust assay that tightly controls biological as well as biophysical conditions and is operated using a standard syringe pump and brightfield/fluorescence microscopy. Parameters such as microcirculatory hemodynamic conditions, endothelial cell type, blood cell type(s) and concentration(s), drug/inhibitory concentration etc., can all be easily controlled. As such, our microsystem provides a method to quantitatively investigate disease processes in which microvascular flow is impaired due to alterations in cell adhesion, aggregation, and deformability, a capability unavailable with existing assays.

A method to culture an endothelial cell monolayer throughout the entire inner 3D surface of a microfluidic device with microvascular-sized channels (&lt;30 &mu;m) is described. This in vitro microvasculature model enables the study of biophysical interactions between blood cells, endothelial cells, and soluble factors in hematologic diseases.

Advances in  microfabrication techniques have enabled the production of inexpensive and  reproducible microfluidic systems for conducting biological and ...

Bioengineering

Assessing Teratogenic Changes in a Zebrafish Model of Fetal Alcohol Exposure

Fetal alcohol syndrome (FAS) is a severe manifestation of embryonic exposure to ethanol. It presents with characteristic defects to the face and organs, including mental retardation due to disordered and damaged brain development. Fetal alcohol spectrum disorder (FASD) is a term used to cover a continuum of birth defects that occur due to maternal alcohol consumption, and occurs in approximately 4% of children born in the United States. With 50% of child-bearing age women reporting consumption of alcohol, and half of all pregnancies being unplanned, unintentional exposure is a continuing issue2. In order to best understand the damage produced by ethanol, plus produce a model with which to test potential interventions, we developed a model of developmental ethanol exposure using the zebrafish embryo. Zebrafish are ideal for this kind of teratogen study3-8. Each pair lays hundreds of eggs, which can then be collected without harming the adult fish. The zebrafish embryo is transparent and can be readily imaged with any number of stains. Analysis of these embryos after exposure to ethanol at different doses and times of duration and application shows that the gross developmental defects produced by ethanol are consistent with the human birth defect. Described here are the basic techniques used to study and manipulate the zebrafish FAS model.

In order to understand the molecular mechanisms of the ethanol-induced developmental damage, we have developed a zebrafish model of ethanol exposure and are exploring the physical, cellular, and genetic alterations that occur after ethanol exposure1. We then seek to find potential interventions and rapidly test them in this animal model.

Fetal alcohol syndrome (FAS) is a severe manifestation of embryonic exposure to ethanol. It presents with characteristic defects to the face and organs, ...

Platelet Adhesion and Aggregation Under Flow using Microfluidic Flow Cells

Platelet aggregation occurs in response to vascular injury where the extracellular matrix below the endothelium has been exposed. The platelet adhesion cascade takes place in the presence of shear flow, a factor not accounted for in conventional (static) well-plate assays. This article reports on a platelet-aggregation assay utilizing a microfluidic well-plate format to emulate physiological shear flow conditions. Extracellular proteins, collagen I or von Willebrand factor are deposited within the microfluidic channel using active perfusion with a pneumatic pump. The matrix proteins are then washed with buffer and blocked to prepare the microfluidic channel for platelet interactions. Whole blood labeled with fluorescent dye is perfused through the channel at various flow rates in order to achieve platelet activation and aggregation. Inhibitors of platelet aggregation can be added prior to the flow cell experiment to generate IC50 dose response data.

The platelet adhesion cascade takes place in the presence of shear flow, a factor not accounted for in conventional (static) well-plate assays. This article reports on a platelet-aggregation assay utilizing a microfluidic well-plate format to emulate physiological shear flow conditions.

Platelet aggregation occurs in response to vascular injury where the extracellular matrix below the endothelium has been exposed. The platelet adhesion ...

Biology

Microfluidic Flow Chambers Using Reconstituted Blood to Model Hemostasis and Platelet Transfusion In Vitro

Blood platelets prepared for transfusion gradually lose hemostatic function during storage. Platelet function can be investigated using a variety of (indirect) in vitro experiments, but none of these is as comprehensive as microfluidic flow chambers. In this protocol, the reconstitution of thrombocytopenic fresh blood with stored blood bank platelets is used to simulate platelet transfusion. Next, the reconstituted sample is perfused in microfluidic flow chambers which mimic hemostasis on exposed subendothelial matrix proteins. Effects of blood donation, transport, component separation, storage and pathogen inactivation can be measured in paired experimental designs. This allows reliable comparison of the impact every manipulation in blood component preparation has on hemostasis. Our results demonstrate the impact of temperature cycling, shear rates, platelet concentration and storage duration on platelet function. In conclusion, this protocol analyzes the function of blood bank platelets and this ultimately aids in optimization of the processing chain including phlebotomy, transport, component preparation, storage and transfusion.

Microfluidic Flow Chambers Using Reconstituted Blood to Model Hemostasis and Platelet Transfusion In Vitro

Platelet transfusion and hemostasis was modeled using blood reconstitution and microfluidic flow chambers to investigate the function of blood banking platelets. The data demonstrate the consequences of platelet storage lesion on hemostasis, in vitro.

Blood platelets prepared for transfusion gradually lose hemostatic function during storage. Platelet function can be investigated using a variety of ...

A Microfluidic Flow Chamber Model for Platelet Transfusion and Hemostasis Measures Platelet Deposition and Fibrin Formation in Real-time

Microfluidic models of hemostasis assess platelet function under conditions of hydrodynamic shear, but in the presence of anticoagulants, this analysis is restricted to platelet deposition only. The intricate relationship between Ca2+-dependent coagulation and platelet function requires careful and controlled recalcification of blood prior to analysis. Our setup uses a Y-shaped mixing channel, which supplies concentrated Ca2+/Mg2+ buffer to flowing blood just prior to perfusion, enabling rapid recalcification without sample stasis. A ten-fold difference in flow velocity between both reservoirs minimizes dilution. The recalcified blood is then perfused in a collagen-coated analysis chamber, and differential labeling permits real-time imaging of both platelet and fibrin deposition using fluorescence video microscopy. The system uses only commercially available tools, increasing the chances of standardization. Reconstitution of thrombocytopenic blood with platelets from banked concentrates furthermore models platelet transfusion, proving its use in this research domain. Exemplary data demonstrated that coagulation onset and fibrin deposition were linearly dependent on the platelet concentration, confirming the relationship between primary and secondary hemostasis in our model. In a timeframe of 16 perfusion min, contact activation did not take place, despite recalcification to normal Ca2+ and Mg2+ levels. When coagulation factor XIIa was inhibited by corn trypsin inhibitor, this time frame was even longer, indicating a considerable dynamic range in which the changes in the procoagulant nature of the platelets can be assessed. Co-immobilization of tissue factor with collagen significantly reduced the time to onset of coagulation, but not its rate. The option to study the tissue factor and/or the contact pathway increases the versatility and utility of the assay.

This paper describes an experimental model of hemostasis that simultaneously measures platelet function and coagulation. Platelet and fibrin fluorescence is measured in real-time, and platelet adhesion rate, coagulation rate, and onset of coagulation are determined. The model is used to determine platelet procoagulant properties under flow in concentrates for transfusion.

Microfluidic models of hemostasis assess platelet function under conditions of hydrodynamic shear, but in the presence of anticoagulants, this analysis is ...

Turbidimetry on Human Washed Platelets: The Effect of the Pannexin1-inhibitor Brilliant Blue FCF on Collagen-induced Aggregation

Turbidimetry is a laboratory technique that is applied to measure the aggregation of platelets suspended in either plasma (platelet-rich plasma, PRP) or in buffer (washed platelets), by the use of one or a combination of agonists. The use of washed platelets separated from their plasma environment and in the absence of anticoagulants allows for studying intrinsic platelet properties. Among the large panel of agonists, arachidonic acid (AA), adenosine di-phosphate (ADP), thrombin and collagen are the most frequently used. The aggregation response is quantified by measuring the relative optical density (OD) over time of platelet suspension under continuous stirring. Platelets in homogeneous suspension limit the passage of light after the addition of an agonist, platelet shape change occurs producing a small transitory increase in OD. Following this initial activation step, platelet clots form gradually, allowing the passage of light through the suspension as a result of decreased OD. The aggregation process is ultimately expressed as a percentage, compared to the OD of platelet-poor plasma or buffer. Rigorous calibration is thus essential at the beginning of each experiment. As a general rule: calibration to 0% is set by measuring the OD of a non-stimulated platelet suspension while measuring the OD of the suspension medium containing no platelets represents a value of 100%. Platelet aggregation is generally visualized as a real-time aggregation curve. Turbidimetry is one of the most commonly used laboratory techniques for the investigation of platelet function and is considered as the historical gold standard and used for the development of new pharmaceutical agents aimed at inhibiting platelet aggregation. Here, we describe detailed protocols for 1) preparation of human washed platelets and 2) turbidimetric analysis of collagen-induced aggregation of human washed platelets pretreated with the food dye Brilliant Blue FCF that was recently identified as an inhibitor of Pannexin1 (Panx1) channels.

We describe a straightforward method for the isolation of washed platelets from human blood followed by agonist-induced platelet aggregation measurements by turbidimetry. As an example we apply this method for studying the aggregation response of human platelets to collagen after a pre-incubation with the Pannexin1 channel inhibitor Brilliant Blue FCF.

Turbidimetry is a laboratory technique that is applied to measure the aggregation of platelets suspended in either plasma (platelet-rich plasma, PRP) or ...

Live-cell Imaging of Platelet Degranulation and Secretion Under Flow

Blood platelets are essential players in hemostasis, the formation of thrombi to seal vascular breaches. They are also involved in thrombosis, the formation of thrombi that occlude the vasculature and injure organs, with life-threatening consequences. This motivates scientific research on platelet function and the development of methods to track cell-biological processes as they occur under flow conditions.
A variety of flow models are available for the study of platelet adhesion and aggregation, two key phenomena in platelet biology. This work describes a method to study real-time platelet degranulation under flow during activation. The method makes use of a flow chamber coupled to a syringe-pump setup that is placed under a wide-field, inverted, LED-based fluorescence microscope. The setup described here allows for the simultaneous excitation of multiple fluorophores that are delivered by fluorescently labeled antibodies or fluorescent dyes. After live-cell imaging experiments, the cover glasses can be further processed and analyzed using static microscopy (i.e., confocal microscopy or scanning electron microscopy).

This work describes a fluorescence microscopy-based method for the study of platelet adhesion, spreading, and secretion under flow. This versatile platform enables the investigation of platelet function for mechanistic research on thrombosis and hemostasis.

Blood platelets are essential players in hemostasis, the formation of thrombi to seal vascular breaches. They are also involved in thrombosis, the ...

Analyzing Platelet Phosphatidylserine and Microvesicle Release

Procoagulant Platelet Characterization by Measuring Phosphatidylserine Exposure and Microvesicle Release from Human Purified Platelets

Activated platelets promote coagulation primarily by exposing the procoagulant phospholipid phosphatidylserine (PS) on their outer membrane surfaces and releasing PS-expressing microvesicles that retain the original membrane architecture and cytoplasmic components of their originating cells. The accessibility of phosphatidylserine facilitates the binding of major coagulation factors, significantly amplifying the catalytic efficiency of coagulation enzymes, while microvesicle release acts as a pivotal mediator of intercellular signaling. Procoagulant platelets play a crucial role in clot stabilization during hemostasis, and their increased proportion in the bloodstream correlates with an increased risk of thrombosis. It has also been shown that platelet microvesicles are rich in growth factors that promote wound healing and inflammatory modulation. Analyzing phosphatidylserine exposure and microvesicle release using flow cytometry poses significant challenges due to their small size and the limited number of positive events for markers of interest. Despite considerable advances in the last decade, methods for assessing phosphatidylserine exposure and microvesicle release remain a work in progress. Unfortunately, no single universally applicable protocol exists, and several factors must be evaluated to determine the most appropriate methodology for each specific application. Here, we describe a detailed protocol for isolating washed platelets from human blood, followed by collagen and/or thrombin activation, to measure the exposure of phosphatidylserine and microvesicle release that characterize procoagulant platelets. This protocol is designed to facilitate the initial preparation of platelet-rich plasma and the isolation of washed platelets. Finally, phosphatidylserine exposure and microvesicle release are quantified by flow cytometry, enabling the identification of procoagulant platelets.

Procoagulant platelet formation has been correlated with an increased risk of thrombosis. Presented here is a precise protocol for isolating washed platelets from human blood, intended to quantify the exposure of phosphatidylserine and microvesicle release, which are distinctive features of procoagulant platelets.

Activated platelets promote coagulation primarily by exposing the procoagulant phospholipid phosphatidylserine (PS) on their outer membrane surfaces and ...

Microfluidics in Assessing Platelet Function

Summary

Explore More Videos

Chapters in this video

Assessing Endothelial Vasodilator Function with the Endo-PAT 2000

In Vivo Canine Muscle Function Assay

Endothelialized Microfluidics for Studying Microvascular Interactions in Hematologic Diseases

Assessing Teratogenic Changes in a Zebrafish Model of Fetal Alcohol Exposure

Platelet Adhesion and Aggregation Under Flow using Microfluidic Flow Cells

Microfluidic Flow Chambers Using Reconstituted Blood to Model Hemostasis and Platelet Transfusion In Vitro

A Microfluidic Flow Chamber Model for Platelet Transfusion and Hemostasis Measures Platelet Deposition and Fibrin Formation in Real-time

Turbidimetry on Human Washed Platelets: The Effect of the Pannexin1-inhibitor Brilliant Blue FCF on Collagen-induced Aggregation

Live-cell Imaging of Platelet Degranulation and Secretion Under Flow

Procoagulant Platelet Characterization by Measuring Phosphatidylserine Exposure and Microvesicle Release from Human Purified Platelets