This work was supported by the National Heart Lung and Blood Institute (NHLBI) of the National Institutes of Health under award numbers R01 HL121212 (PI: Sen Gupta), R01 HL129179 (PI: Sen Gupta, Co-I: Li) and R01 HL098217 (PI: Nieman). The content of this publication is solely the responsibility of the authors and does not necessarily represent the official views of the National Institutes of Health.

All procedures and manipulations of animals have been approved by Institutional Animal Care and Use Committees (IACUC) of The Cleveland Clinic in accordance with the United States Public Health Service Policy on the Humane Care and Use of Animals, and the NIH Guide for the Care and Use of Laboratory Animals.
1. Preparations:
<ol>
	<li>Fluorescent Dye for Labeling Platelets
	<ol>
		<li>Prepare rhodamine 6G solution, 0.5 mg/ml, in saline and sterilize the solution with 0.22 &#181;m filter.</li>
	</ol>
	</li>
	<li>FeCl3 Solution
	<ol>
		<li>Make a fresh stock solution of 30% FeCl3 (Anhydrous, equals 1.85 M) in pure water, and filter with 0.45 &#181;m filter. Prepare 5 ml fresh FeCl3 solution at the desired concentration [e.g., 2.5% (0.154 M), 5% (0.308 M), 7.5% (0.463 M), 10% (0.617 M) and 12.5% (0.771 M)] by diluting the 30% FeCl3 solution with water in a 6 cm tissue culture dish and keep the lid closed. 
		NOTE: Using the culture dish makes it easier to pick up the filter paper saturated with FeCl3 solution than to pick it up from a narrow container, such as a 1.5 ml microcentrifuge tube.&#160;&#160;FeCl3 is extremely hazardous and that precautions, such as wearing gloves and lab coat etc., Personal Protective Equipment, should always be taken.</li>
	</ol>
	</li>
	<li>Filter Papers
	<ol>
		<li>Cut the filter paper to 1 mm x 2 mm size. Soak enough pieces of the cut filter paper in the FeCl3 solution in the same dish.</li>
	</ol>
	</li>
	<li>Papaverine Hydrochloride Solution
	<ol>
		<li>Prepare papaverine hydrochloride solution (25 mg/ml) in water and sterilize the solution with 0.22 &#181;m filter.</li>
	</ol>
	</li>
	<li>Agarose Gel
	<ol>
		<li>Prepare 2% agarose gel in PBS or saline in 6-well tissue culture plate, ~ 1 cm thickness, and cut it to half-circle with ~ 3 cm diameter.</li>
	</ol>
	</li>
</ol>
2. FeCl3 Induced Carotid Arterial Injury Thrombosis Model
<ol>
	<li>Surgical Procedures for the Thrombosis Model
	<ol>
		<li>Use 8 - 12 weeks old C57Bl6 mice (or other strains as necessary). Anesthetize mice with the mixture of ketamine (100 mg/kg)/xylazine (10 mg/kg) via intraperitoneal injection. Confirm the depth of the anesthesia by toe pinching. 
		NOTE: This amount of ketamine/xylazine is sufficient to keep the animal in stable anesthesia and no pain (response to toe pinching) for at least 1 hr.</li>
		<li>Remove fur on the neck and upper chest with a small animal electric clipper. Depilatory cream is not necessary. Apply neutral petroleum eye&#160;ointment on the eyes to prevent dryness during anesthesia.</li>
		<li>Secure the mouse in the supine position on the lid of 15 cm tissue culture plate. Use one long tape (~ 10 cm) to secure hind limbs and lower body, and two pieces of small tape (2 cm x 0.5 cm) to secure forelegs. Use a 4-0 suture to wrap around the incisors, tape the two ends of the suture to the lid to keep mouse neck straight (Figure 2).</li>
		<li>Place plate with mouse head toward the operator under a surgical light source.</li>
		<li>Sterilize the surgical site with alcohol pad. Use the Micro-Adson Forceps to hold the skin and use a surgical scissors to make a small incision (2 - 3 mm).</li>
		<li>Hold the skin of the incision with the forceps and insert surgical scissors with jaws closed into the incision. Push the scissors subcutaneously toward the manubrium or chin, and then open the scissors to free the skin from the subcutaneous layer.</li>
		<li>Cut the skin with the surgical scissors to make a midline incision from the manubrium to the level of hyoid bone (Figure 3A and 3B). Bluntly dissect the thin fascia (dotted line, in Figure 3B) between the submaxillary glands with a fine hemostat and a Graefe forceps (Figure 3B and 3C; blue arrows). 
		NOTE: The manubrium (black arrow in Figure 3C) and the trachea (T in Figure 3C) will be seen after this step.</li>
		<li>Hold the soft tissue and the skin seen at the right of the manubrium with the Graefe forceps, insert the hemostat under the fascia toward the 2 o&#39;clock position (yellow dotted line in Figure 3C), open the jaws of the hemostat to free the jugular vein from surrounding tissue.</li>
		<li>Cut the fascia, soft tissue and skin toward the 2 o&#39;clock position (Figure 3C) with the surgical scissors to expose the right jugular vein (Figure 3D, arrow).</li>
		<li>Draw about 200 - 300 &#181;l rhodamine 6G solution using a 1 ml syringe with 22 G needle, and then change it to 30 G needle. 
		NOTE: This protects the 30 G needle from damage of its fine tip.&#160; Directly drawing the rhodamine 6G solution with the 30 G needle will not damage the tip; however touching the container wall accidentally will cause damage, which may make injection difficult.</li>
		<li>Flush the 30 G needle by slowly pushing out the rhodamine 6G solution, make sure no air bubbles remain in the syringe or the needle. Keep 100 &#181;l rhodamine 6G solution for injection.</li>
		<li>Bend 2 - 3 mm tip of the needle to a 90&#176; angle with the needle holder, which will prevent inserting the needle too deep to the jugular vein and makes the injection more easily controlled (Figure 3D).</li>
		<li>Inject the rhodamine 6G solution into the right jugular vein to label platelets. To stabilize the syringe and keep the needle in position, hold the syringe with one hand and the needle with the Graefe forceps with another hand during the injection.After injection, clamp the injection site with the Graefe forceps to avoid bleeding.</li>
		<li>Open the jaws of the hemostat and insert it under the Graefe forceps to clamp the vessel wall of the injection site, and then ligate the injection site with a 6-0 suture. 
		NOTE: As an alternative of step 2.1.15, applying pressure to the injection site with a finger for 2 min will stop the bleeding; however, this method has a risk of causing further bleeding if the clot at the injection site is removed accidently.</li>
		<li>Bluntly dissect the soft tissue and fascia around the left submaxillary gland with the hemostat and the Graefe forceps and pull the gland toward the 7 o&#39;clock position (Figure 3E) to expose the left sternocleidomastoid muscle (blue arrow in Figure 3E).</li>
		<li>Bluntly dissect the fascia (Figure 3E, dotted line) between the left sternocleidomastoid muscle and the omohyoid muscle or the sternohyoid muscle (Figure 3E, green arrow) located to the left site of the trachea with the hemostat.</li>
		<li>Pass a needle with 6-0 silk suture (about 15 cm long) under the middle of the sternocleidomastoid muscle (blue arrow in Figure 3E), put the two ends of the suture together and pull the suture laterally toward the 10 o&#39;clock position. 
		NOTE: This procedure should be performed carefully to avoid injury of the left jugular vein, which is located outside of the sternocleidomastoid muscle.</li>
		<li>Bluntly separate the thin sternohyoid muscle and/or omohyoid muscle to expose the carotid artery (CA) (Figure 3F arrow) after pulling away the sternocleidomastoid muscle. Cut the sternohyoid muscle and/or omohyoid muscle as necessary. Use the hemostat to separate the soft tissue from CA without touching the CA. 
		NOTE: CA is accompanied by the vagus nerve, the white structure seen in Figure 3G (green arrow), and in most cases it is under the thin sternohyoid muscle and/or omohyoid muscle (between the yellow dotted lines in Figure 3F).</li>
		<li>Use a fine tip forceps to pick up the lateral fascia around the CA while avoiding the vagus nerve and CA. Use another fine tip forceps to punch a hole on the fascia between the CA and vagus nerve.</li>
		<li>Pass the hook through the hole and gently lift the CA, and then place the fine tip forceps under the CA. Move the hook and forceps in opposite directions along the CA to strip adventitial soft tissue. Free at least ~ 5 mm length of CA (Figure 3H, blue arrow) from surrounding tissue.</li>
		<li>To block background fluorescence, press the end of a black plastic coffee stirrer to flat, crosscut the coffee stirrer into a 3 mm piece, and then cut longitudinally along the fold edges to get two pieces of &#34;U&#34; shape plastic (Figure 3H, green arrow).</li>
		<li>Rinse one piece of this &#34;U&#34; shape plastic with saline and hold it with forceps and place it under the CA while gently lifting the CA with the hook (Figure 3H, green arrow). Immediately apply 2-3 drops of saline to avoid drying the CA.</li>
	</ol>
	</li>
	<li>Real time Recording Video
	<ol>
		<li>Transfer the mouse and plate lid together to the microscope stage and place in an appropriate position under the 10X water lens. Fill the space between the 10X lens and the CA with saline.</li>
		<li>Start the digital video recording software application, and launch a new file and name it accordingly. Record normal vessel images and write down the frame number when the video recording is stopped. 
		NOTE: Reference the video recording software in the computer for recording. We use digital video recording software and the following descriptions are based on this application.&#160; 
		NOTE: Since mechanical trauma can injure the endothelium and lead to thrombus formation20, it is necessary to confirm that there is no surgical injury to the vessel wall prior to the FeCl3 injury. It is also necessary to confirm that there is no remaining tissue around the CA, which may form a barrier and attenuate the effect of FeCl3-induced injury. Write down the frame number of the records will make it easy to analyze the videos according elapsed times later.</li>
		<li>Move the mouse with plate lid out of the 10X lens. Fold a corner of a paper towel to make a thin and fine tip and use it to carefully blot the saline around the CA (avoid touching CA) as well as in other places in the surgical field. This is important to avoid dilution of the FeCl3 solution used.</li>
		<li>Use a fine tip forceps and pick up a piece of filter paper saturated with FeCl3 solution and place it directly on the CA and keep it on site for 1 min. 
		NOTE: To aid visualization of the bloodstream and harvest accurate data, place the filter paper close to the distal end of the exposed CA (Figure 3I) and leave a short segment at the proximal site (green arrow in Figure 3I) for observing blood flow at late phase (see below).</li>
		<li>Remove the filter paper (this time point is defined as the start of &#8220;after injury&#8221;) and rinse the CA with saline (at least 2 ml). &#160;</li>
		<li>Put the mouse with plate lid back to the 10X lens; observe the vessel and start to record video images. Write down the video frame number at the end of the first minute of recording. 
		NOTE: Thrombus formation is identified by accumulation of the fluorescent platelets, which is observed in real-time video images on computer screen or under microscope. Record video images immediately after putting the mouse back under the microscope. Keep recording to the end of the first minute after removing the filter paper. Observe the entire vessel from the distal to the proximal sites to obtain information about the injury, and then focus on the area of interest (usually is the center of the injured area) for further imaging.</li>
		<li>Record video images for 10 sec every minute for the first 10 min (e.g., 2:55 to 3:05) and then 10 sec every other minute until the end of the experiment. Write down the frame numbers when each recording is stopped. 
		NOTE: The end points of the model are: 1) when blood flow has ceased for &#62;&#160; 30 sec; or 2) if occlusion is not seen in 30 min after injury. In this case, use 30 min for statistical analysis. Set Exposure Time of the video record as 10 msec at the beginning, so it will be easy to observe the blood flow over the thrombi. When the thrombi become large and the fluorescence saturates the camera&#8217;s sensor, it becomes difficult to identify the blood flow over thrombi. To solve this issue, move the imaging center to the proximal uninjured site (Figure 3I, green arrow) where blood flow can still be clearly observed. We also increase the Expose Time to 15 or 20 msec when focusing on the proximal un-injured site, so the blood flow can be observed more clearly. When blood flow will cease, numerous larger cells (leukocytes) start to roll on the vessel wall at the proximal site of the thrombus. Flow usually stops within 2 - 3 min after the appearance of the large cells. Write down the Expose Time on the recording sheet if it is changed. It usually takes about 30 min from the anesthesia to the end of the experiment, if the normal wild type C57Bl6 mice were used. 
		CAUTION NOTE: Don&#39;t use the white aggregated platelet clot as the sign of the blood flow cessation, which will not give an accurate data and it is affected by the exposure time. The white clot covered the vessel lumen does not mean blood flow ceased (see the representative video 1).</li>
		<li>At the end of experiment, euthanize the mice using overdose ketamine/xylazine (200/20 mg/Kg), followed by cervical dislocation after confirming no breath and heartbeat.</li>
	</ol>
	</li>
</ol>
3. FeCl3 Induced Mesenteric Artery/Vein Thrombosis Model
<ol>
	<li>Anesthetize 8 &#8722; 12 week old C57Bl6 mice as mentioned in section 2.1.1. Apply vet ointment on the eyes to prevent dryness during anesthesia.</li>
	<li>Inject Papaverine solution (total 0.4 mg/mouse) intraperitoneally to inhibit gut peristalsis.</li>
	<li>Remove fur on the neck and abdomen with an animal electric clipper. Depilatory cream is not necessary.</li>
	<li>Secure the mouse in the supine position on the lid of 15 cm tissue culture plate. Use four pieces of small tape (2 cm x 0.5 cm) to secure all legs. Use a 4 -0 suture to wrap around the incisors, tape the two ends of the suture to the lid to keep the neck straight (Figure 2).</li>
	<li>Follow procedures 2.1.4 - 2.1.15 to expose the jugular vein for injection of rhodamine 6G to label platelets.</li>
	<li>Perform a midline incision through the skin from xiphoid to lower abdomen using the surgical scissors (Figure 4A). Pick up the middle peritoneum (also called linea alba, Figure 4A, arrow) with Graefe forceps, which is clear and has no blood vessels, lift about 2 - 3 mm high, confirm no bowel under the cutting line, and cut the peritoneum open longitudinally with the fine scissors (Figure 4B).</li>
	<li>Re-secure the mice to right lateral position. Place the half-circle agarose gel close to the abdomen, and gently exteriorize the intestines and spread it on the top of the gel with the Graefe forceps (Figure 4C). Use the second branches for the thrombosis experiment (Figure 4C, arrow). 
	NOTE: Use the hemostat or Micro-Adson forceps to help to exteriorize the intestines. Avoid hurting the bowel and the vessel.</li>
	<li>Place 5 - 6 drops saline to keep the moisture of the bowel and vessel. Transfer the mouse with the plate lid together to the microscope stage and place in appropriate position under the 10X dry lens. 
	NOTE: Use a dry lens because the jejunoileal membrane cannot hold saline solution. Make sure to drop saline solution on the exposed tissues to keep them moist.</li>
	<li>Blot the saline around the mesenteric artery and vein before treatment.</li>
	<li>Use a fine tip forceps to pick up a piece of filter paper saturated with 12.5% FeCl3 solution and place it directly on the mesenteric artery and vein for 1 min (Figure 4D).&#160; Remove the filter paper (this time point is defined as the start of &#8220;after injury&#8221;) and rinse the injured mesenteric artery and vein with saline (at least 2 ml). &#160;</li>
	<li>Put the mouse with plate lid back under the 10X dry lens; observe the vessels and start to record video images as mentioned in 2.2). 
	NOTE: The end points of this experiment are: 1) when blood flow has ceased for &#62; 30 sec; or 2) if occlusion is not seen in 30 min after injury, and in this case use 30 min for statistical analysis.</li>
	<li>At the end of experiment, euthanize the mice using overdose ketamine/xylazine (200/20 mg/Kg), followed by cervical dislocation after no breath and heart beating are confirmed.</li>
</ol>

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C., et al. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Platelet-like+nanoparticles:+mimicking+shape,+flexibility,+and+surface+biology+of+platelets+to+target+vascular+injuries.">Platelet-like nanoparticles: mimicking shape, flexibility, and surface biology of platelets to target vascular injuries.</a> ACS Nano. 8, 11243-11253 (2014).</li><li>Modery, C. L., et al. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Heteromultivalent+liposomal+nanoconstructs+for+enhanced+targeting+and+shear-stable+binding+to+active+platelets+for+site-selective+vascular+drug+delivery.">Heteromultivalent liposomal nanoconstructs for enhanced targeting and shear-stable binding to active platelets for site-selective vascular drug delivery.</a> Biomaterials. 32, 9504-9514 (2011).</li><li>Woollard, K. J., Sturgeon, S., Chin-Dusting, J. P., Salem, H. H., Jackson, S. P. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Erythrocyte+hemolysis+and+hemoglobin+oxidation+promote+ferric+chloride-induced+vascular+injury.">Erythrocyte hemolysis and hemoglobin oxidation promote ferric chloride-induced vascular injury.</a> J Biol Chem. 284, 13110-13118 (2009).</li><li>Ciciliano, J. C., et al. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Resolving+the+multifaceted+mechanisms+of+the+ferric+chloride+thrombosis+model+using+an+interdisciplinary+microfluidic+approach.">Resolving the multifaceted mechanisms of the ferric chloride thrombosis model using an interdisciplinary microfluidic approach.</a> Blood. 126, 817-824 (2015).</li><li>Barr, J. D., Chauhan, A. K., Schaeffer, G. V., Hansen, J. K., Motto, D. G. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Red+blood+cells+mediate+the+onset+of+thrombosis+in+the+ferric+chloride+murine+model.">Red blood cells mediate the onset of thrombosis in the ferric chloride murine model.</a> Blood. 121, 3733-3741 (2013).</li><li>Dunne, E., et al. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Cadherin+6+has+a+functional+role+in+platelet+aggregation+and+thrombus+formation.">Cadherin 6 has a functional role in platelet aggregation and thrombus formation.</a> Arterioscler Thromb Vasc Biol. 32, 1724-1731 (2012).</li><li>Lockyer, S., et al. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=GPVI-deficient+mice+lack+collagen+responses+and+are+protected+against+experimentally+induced+pulmonary+thromboembolism.">GPVI-deficient mice lack collagen responses and are protected against experimentally induced pulmonary thromboembolism.</a> Thromb Res. 118, 371-380 (2006).</li><li>Zhou, J., et al. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=The+C-terminal+CGHC+motif+of+protein+disulfide+isomerase+supports+thrombosis.">The C-terminal CGHC motif of protein disulfide isomerase supports thrombosis.</a> J Clin Invest. , (2015).</li><li>Eckly, A., et al. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Mechanisms+underlying+FeCl3-induced+arterial+thrombosis.">Mechanisms underlying FeCl3-induced arterial thrombosis.</a> J Thromb Haemost. 9, 779-789 (2011).</li><li>Day, S. M., Reeve, J. L., Myers, D. D., Fay, W. P. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Murine+thrombosis+models.">Murine thrombosis models.</a> Thromb Haemost. 92, 486-494 (2004).</li><li>Cooley, B. C. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Murine+models+of+thrombosis.">Murine models of thrombosis.</a> Thromb Res. 129 Suppl 2, S62-S64 (2012).</li><li>Gupta, N., Li, W., McIntyre, T. M. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Deubiquitinases+Modulate+Platelet+Proteome+Ubiquitination,+Aggregation,+and+Thrombosis.">Deubiquitinases Modulate Platelet Proteome Ubiquitination, Aggregation, and Thrombosis.</a> Arterioscler Thromb Vasc Biol. 35, 2657-2666 (2015).</li><li>Konstantinides, S., et al. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Distinct+antithrombotic+consequences+of+platelet+glycoprotein+Ibalpha+and+VI+deficiency+in+a+mouse+model+of+arterial+thrombosis.">Distinct antithrombotic consequences of platelet glycoprotein Ibalpha and VI deficiency in a mouse model of arterial thrombosis.</a> J Thromb Haemost. 4, 2014-2021 (2006).</li><li>Versteeg, H. H., Heemskerk, J. W., Levi, M., Reitsma, P. H. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=New+fundamentals+in+hemostasis.">New fundamentals in hemostasis.</a> Physiol Rev. 93, 327-358 (2013).</li><li>Yan, S. F., Mackman, N., Kisiel, W., Stern, D. M., Pinsky, D. J. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Hypoxia/Hypoxemia-Induced+activation+of+the+procoagulant+pathways+and+the+pathogenesis+of+ischemia-associated+thrombosis.">Hypoxia/Hypoxemia-Induced activation of the procoagulant pathways and the pathogenesis of ischemia-associated thrombosis.</a> Arterioscler Thromb Vasc Biol. 19, 2029-2035 (1999).</li><li>Rahaman, S. O., Li, W., Silverstein, R. L. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Vav+Guanine+nucleotide+exchange+factors+regulate+atherosclerotic+lesion+development+in+mice.">Vav Guanine nucleotide exchange factors regulate atherosclerotic lesion development in mice.</a> Arterioscler Thromb Vasc Biol. 33, 2053-2057 (2013).</li><li>Silverstein, R. L., Li, W., Park, Y. M., Rahaman, S. O. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Mechanisms+of+cell+signaling+by+the+scavenger+receptor+CD36:+implications+in+atherosclerosis+and+thrombosis.">Mechanisms of cell signaling by the scavenger receptor CD36: implications in atherosclerosis and thrombosis.</a> Trans Am Clin Climatol Assoc. 121, 206-220 (2010).</li><li>Liu, J., Li, W., Chen, R., McIntyre, T. M. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Circulating+biologically+active+oxidized+phospholipids+show+on-going+and+increased+oxidative+stress+in+older+male+mice.">Circulating biologically active oxidized phospholipids show on-going and increased oxidative stress in older male mice.</a> Redox Biol. 1, 110-114 (2013).</li></ol>

The authors have nothing to disclose.

The FeCl3-induced model is one of the most widely used thrombosis models, which can not only provide valuable information about genetic modifications on platelet function and thrombosis7,8,16,19,31-33, but can also be a valuable tool for evaluation of therapeutic compounds and strategies for treatment and prevention of atherothrombotic diseases11,17,34-37. Here we have shown our modifications and refinements of this model and showed additional evidence of the utility of this technique, wh...

Arterial thrombosis (blood clot) is a common complication of many systemic diseases associated with chronic inflammation, including atherosclerosis, diabetes, obesity, cancer and chronic autoimmune rheumatologic disorders. Thrombi that occur in the arterial circulation stem from inappropriate platelet activation, aggregation and subsequent coagulatory mechanisms, and are implicated in heart attacks, strokes and extremity loss. The vessel wall is a complex system that includes multiple cell types and is influenced by a multitude of extrinsic factors including shear stress, circulating blood cells, hormones and cytokines, as well as expression of antioxidant proteins in the vessel wall. In vitro experiments cannot replicate this complex environment and therefore in vivo studies using animal models are critical to allow better understanding of mechanisms involved in thrombotic disorders.
Mice have been shown to have similar mechanisms to humans in terms of thrombosis, atherosclerosis, inflammation and diabetes1,2. Furthermore, transgenic and knockout mice can be created to test the function of specific gene products in a complex physiologic or pathologic environment. Such studies mimic human pathology and may provide important mechanistic information related to discovery of new pathways and therapies, as well as provide important details in characterizing drug effects on thrombosis.
Pathological arterial thrombi occur due to endothelial layer injury or dysfunction and exposure of the blood stream to the subendothelial matrix3,4. Various thrombosis models have been developed to induce this endothelial damage such as mechanical injury, photoreactive compound Rose Bengal-based oxidative injury and laser injury5. In this spectrum, Ferric chloride (FeCl3)-induced vascular injury is a widely used model of thrombosis. This reagent when applied to the outer aspect of vessels induces oxidative damage to vascular cells6-8, with loss of endothelial cell protection from circulating platelets and components of the coagulation cascade. The FeCl3 model is simple and sensitive to both anticoagulant and anti-platelets drugs, and has been performed on carotid and femoral arteries, jugular veins, and mesenteric and cremasteric arterioles and venules in mice, rats, guinea pigs and rabbits6-15.
One measurable parameter in this model is the elapsed time from injury to complete vessel occlusion, measured as blood flow cessation with a Doppler flow meter or under direct observation with intravital microscopy6,7,9. A range of times between 5 to 30 min has been reported in different studies in C57Bl6 mice7-10,16, suggesting that FeCl3 concentrations, types of anesthesia, surgical techniques, mouse age, genomic background, method of measuring blood flow, and other environmental variables have significant effects in this model. This wide variability makes it difficult to compare studies from different research groups and may make detection of subtle differences difficult.
With a vision to minimize such variabilities and establish a uniformly reproducible in vivo model system, we have refined the FeCl3-induced carotid artery model that makes the data highly reproducible with minimal variation6-10,16-19. In this paper we describe and share the skills and report several representative experimental examples that can benefit from this model.

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			<td>To fix the mouse to the plate</td>
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			<td style="width:325px;">http://www.npielectronic.de/products/micropositioners/burleigh/gibraltar.html</td>
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ferric chloride-induced murine thrombosis models

Arterial thrombosis (blood clot) is a common complication of many systemic diseases associated with chronic inflammation, including atherosclerosis, diabetes, obesity, cancer and chronic autoimmune rheumatologic disorders. Thrombi are the cause of most heart attacks, strokes and extremity loss, making thrombosis an extremely important public health problem. Since these thrombi stem from inappropriate platelet activation and subsequent coagulation, targeting these systems therapeutically has important clinical significance for developing safer treatments. Due to the complexities of the hemostatic system, in vitro experiments cannot replicate the blood-to-vessel wall interactions; therefore, in vivo studies are critical to understand pathological mechanisms of thrombus formation. To this end, various thrombosis models have been developed in mice. Among them, ferric chloride (FeCl3) induced vascular injury is a widely used model of occlusive thrombosis that reports platelet activation and aggregation in the context of an aseptic closed vascular system. This model is based on redox-induced endothelial cell injury, which is simple and sensitive to both anticoagulant and anti-platelets drugs. The time required for the development of a thrombus that occludes blood flow gives a quantitative measure of vascular injury, platelet activation and aggregation that is relevant to thrombotic diseases. We have significantly refined this FeCl3-induced vascular thrombosis model, which makes the data highly reproducible with minimal variation. Here we describe the model and present representative data from several experimental set-ups that demonstrate the utility of this model in thrombosis research.

Carotid Artery Thrombosis Model 
In mice with C57BL6 background, we recommend using 7.5% FeCl3 to treat the vessel for 1 min as a starting point. Under treatment of 7.5% FeCl3, borders of the injured area and normal vessel wall are easily identified under microscope (See online video 1), suggesting that the endothelial layer was significantly damaged. The thrombi formed immediately upon FeCl3 treatment, and are observ...

Watch this Scientific Journal Video about Ferric Chloride-induced Murine Thrombosis Models at JoVE.com

Ferric Chloride-induced Murine Thrombosis Models

We report a refined procedure of the ferric chloride (FeCl3)-induced thrombosis models on carotid and mesenteric artery as well as vein, characterized efficiently using intravital microscopy to monitor time to occlusive thrombi formation.

Arterial thrombosis (blood clot) is a common complication of many systemic diseases associated with chronic inflammation, including atherosclerosis, ...

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21.3K Views.  Lerner Research Institute, Cleveland Clinic. We report a refined procedure of the ferric chloride (FeCl3)-induced thrombosis models on carotid and mesenteric artery as well as vein, characterized efficiently using intravital microscopy to monitor time to occlusive thrombi formation.

Video: Ferric Chloride-induced Murine Thrombosis Models

Electrolytic Inferior Vena Cava Model (EIM) of Venous Thrombosis

Animal models serve a vital role in deep venous thrombosis (DVT) research in order to study thrombus formation, thrombus resolution and to test potential therapeutic compounds (1). New compounds to be utilized in the treatment and prevention of DVT are currently being developed. The delivery of potential therapeutic antagonist compounds to an affected thrombosed vein has been problematic. In the context of therapeutic applications, a model that uses partial stasis and consistently generates thrombi within a major vein has been recently established. The Electrolytic Inferior vena cava Model (EIM) is mouse model of DVT that permits thrombus formation in the presence of continuous blood flow. This model allows therapeutic agents to be in contact with the thrombus in a dynamic fashion, and is more sensitive than other models of DVT (1). In addition, this thrombosis model closely simulates clinical situations of thrombus formation and is ideal to study venous endothelial cell activation, leukocyte migration, venous thrombogenesis, and to test therapeutic applications (1). The EIM model is technically simple, easily reproducible, creates consistent thrombi sizes and allows for a large sample (i.e. thrombus and vein wall) which is required for analytical purposes.

Electrolytic Inferior Vena Cava Model EIM of Venous Thrombosis

The electrolytic induction of endothelial activation to the internal surface of the Inferior Vena Cava results in venous type thrombus formation due to endothelial activation and partial blood stasis, two components of Virchow's triad.

Animal models serve a vital role in deep venous thrombosis (DVT) research in order to study thrombus formation, thrombus resolution and to test potential ...

Mouse Complete Stasis Model of Inferior Vena Cava Thrombosis

Venous thromboembolism (VTE) includes both deep vein thrombosis (DVT) and pulmonary embolism (PE). In the United States (U.S.), the high morbidity and mortality rates make VTE a serious health concern 1-2. After heart disease and stroke, VTE is the third most common vascular disease 3. In the U.S. alone, there is an estimated 900,000 people affected each year, with 300,000 deaths occurring annually 3. A reliable in vivo animal model to study the mechanisms of this disease is necessary.
The advantages of using the mouse complete stasis model of inferior vena cava thrombosis are several. The mouse model allows for the administration of very small volumes of limited availability test agents, reducing costs dramatically. Most promising is the potential for mice with gene knockouts that allow specific inflammatory and coagulation factor functions to be delineated. Current molecular assays allow for the quantitation of vein wall, thrombus, whole blood, and plasma for assays. However, a major concern involving this model is the operative size constraints and the friability of the vessels. Also, due to the small IVC sample weight (mean 0.005 grams) it is necessary to increase animal numbers for accurate statistical analysis for tissue, thrombus, and blood assays such as real-time polymerase chain reaction (RT-PCR), western blot, enzyme-linked immunosorbent (ELISA), zymography, vein wall and thrombus cellular analysis, and whole blood and plasma assays 4-8.
The major disadvantage with the stasis model is that the lack of blood flow inhibits the maximal effect of administered systemic therapeutic agents on the thrombus and vein wall.

The mouse complete stasis model of inferior vena cava thrombosis yields quantifiable amounts of vein wall tissue and thrombus. It has proven useful for evaluating interactions between the vein wall and the occlusive thrombus and in assessing the progression from acute to chronic inflammation.

Venous thromboembolism (VTE) includes both deep vein thrombosis (DVT) and pulmonary embolism (PE).  In the United States (U.S.), the high morbidity and ...

Segmentation and Measurement of Fat Volumes in Murine Obesity Models Using X-ray Computed Tomography

Obesity is associated with increased morbidity and mortality as well as reduced metrics in quality of life.1 Both environmental and genetic factors are associated with obesity, though the precise underlying mechanisms that contribute to the disease are currently being delineated.2,3 Several small animal models of obesity have been developed and are employed in a variety of studies.4 A critical component to these experiments involves the collection of regional and/or total animal fat content data under varied conditions.
Traditional experimental methods available for measuring fat content in small animal models of obesity include invasive (e.g. ex vivo measurement of fat deposits) and non-invasive (e.g. Dual Energy X-ray Absorptiometry (DEXA), or Magnetic Resonance (MR)) protocols, each of which presents relative trade-offs. Current invasive methods for measuring fat content may provide details for organ and region specific fat distribution, but sacrificing the subjects will preclude longitudinal assessments. Conversely, current non-invasive strategies provide limited details for organ and region specific fat distribution, but enable valuable longitudinal assessment. With the advent of dedicated small animal X-ray computed tomography (CT) systems and customized analytical procedures, both organ and region specific analysis of fat distribution and longitudinal profiling may be possible. Recent reports have validated the use of CT for in vivo longitudinal imaging of adiposity in living mice.5,6 Here we provide a modified method that allows for fat/total volume measurement, analysis and visualization utilizing the Carestream Molecular Imaging Albira CT system in conjunction with PMOD and Volview software packages.

Fat content analysis is routinely conducted in studies utilizing murine obesity models. Emerging methods in small animal CT imaging and analysis are providing for longitudinal detail rich fat content analysis. Here we detail step by step procedures for performing small animal CT imaging, analysis, and visualization.

Obesity is associated with increased morbidity and mortality as well as reduced metrics in quality of life.1 Both environmental and genetic factors are ...

Quantitative Analysis and Characterization of Atherosclerotic Lesions in the Murine Aortic Sinus

Atherosclerosis is a disease of the large arteries and a major underlying cause of myocardial infarction and stroke. Several different mouse models have been developed to facilitate the study of the molecular and cellular pathophysiology of this disease. In this manuscript we describe specific techniques for the quantification and characterization of atherosclerotic lesions in the murine aortic sinus and ascending aorta. The advantage of this procedure is that it provides an accurate measurement of the cross-sectional area and total volume of the lesion, which can be used to compare atherosclerotic progression across different treatment groups. This is possible through the use of the valve leaflets as an anatomical landmark, together with careful adjustment of the sectioning angle. We also describe basic staining methods that can be used to begin to characterize atherosclerotic progression. These can be further modified to investigate antigens of specific interest to the researcher. The described techniques are generally applicable to a wide variety of existing and newly created dietary and genetically-induced models of atherogenesis.

We describe procedures to quantify and characterize atherosclerotic lesions in mouse models using precision sectioning of the aortic sinus and ascending aorta combined with histochemical and immunohistochemical analysis.

Atherosclerosis is a disease of the large arteries and a major underlying cause of myocardial infarction and stroke. Several different mouse models have ...

Intrastriatal Injection of Autologous Blood or Clostridial Collagenase as Murine Models of Intracerebral Hemorrhage

Intracerebral hemorrhage (ICH) is a common form of cerebrovascular disease and is associated with significant morbidity and mortality. Lack of effective treatment and failure of large clinical trials aimed at hemostasis and clot removal demonstrate the need for further mechanism-driven investigation of ICH. This research may be performed through the framework provided by preclinical models. Two murine models in popular use include intrastriatal (basal ganglia) injection of either autologous whole blood or clostridial collagenase. Since, each model represents distinctly different pathophysiological features related to ICH, use of a particular model may be selected based on what aspect of the disease is to be studied. For example, autologous blood injection most accurately represents the brain&#39;s response to the presence of intraparenchymal blood, and may most closely replicate lobar hemorrhage. Clostridial collagenase injection most accurately represents the small vessel rupture and hematoma evolution characteristic of deep hemorrhages. Thus, each model results in different hematoma formation, neuroinflammatory response, cerebral edema development, and neurobehavioral outcomes. Robustness of a purported therapeutic intervention can be best assessed using both models. In this protocol, induction of ICH using both models, immediate post-operative demonstration of injury, and early post-operative care techniques are demonstrated. Both models result in reproducible injuries, hematoma volumes, and neurobehavioral deficits. Because of the heterogeneity of human ICH, multiple preclinical models are needed to thoroughly explore pathophysiologic mechanisms and test potential therapeutic strategies.

Preclinical models of intracerebral hemorrhage are utilized to mimic certain aspects of clinical disease. Thus, mechanisms of injury and potential therapeutic strategies may be explored. In this protocol, two models of intracerebral hemorrhage are described, intrastriatal (basal ganglia) injections of autologous blood or collagenase.

Intracerebral hemorrhage (ICH) is a common form of cerebrovascular disease and is associated with significant morbidity and mortality. Lack of effective ...

Orthotopic Injection of Breast Cancer Cells into the Mammary Fat Pad of Mice to Study Tumor Growth.

Breast cancer growth can be studied in mice using a plethora of models. Genetic manipulation of breast cancer cells may provide insights into the functions of proteins involved in oncogenic progression or help to discover new tumor suppressors. In addition, injecting cancer cells into mice with different genotypes might provide a better understanding of the importance of the stromal compartment. Many models may be useful to investigate certain aspects of disease progression but do not recapitulate the entire cancerous process. In contrast, breast cancer cells engraftment to the mammary fat pad of mice better recapitulates the location of the disease and presence of the proper stromal compartment and therefore better mimics human cancerous disease. In this article, we describe how to implant breast cancer cells into mice orthotopically and explain how to collect tissues to analyse the tumor milieu and metastasis to distant organs. Using this model, many aspects (growth, angiogenesis, and metastasis) of cancer can be investigated simply by providing a proper environment for tumor cells to grow.

Cancer is a complex disease that is influenced by the tissue surrounding the tumor as well as local pro- and anti-inflammatory mediators. Therefore, orthotropic injection models, rather than subcutaneous models may be useful to study cancer progression in a manner that better mimics human pathology.

Breast cancer growth can be studied in mice using a plethora of models. Genetic manipulation of breast cancer cells may provide insights into the ...

Adapting Human Videofluoroscopic Swallow Study Methods to Detect and Characterize Dysphagia in Murine Disease Models

This study adapted human videofluoroscopic swallowing study (VFSS) methods for use with murine disease models for the purpose of facilitating translational dysphagia research. Successful outcomes are dependent upon three critical components: test chambers that permit self-feeding while standing unrestrained in a confined space, recipes that mask the aversive taste/odor of commercially-available oral contrast agents, and a step-by-step test protocol that permits quantification of swallow physiology. Elimination of one or more of these components will have a detrimental impact on the study results. Moreover, the energy level capability of the fluoroscopy system will determine which swallow parameters can be investigated. Most research centers have high energy fluoroscopes designed for use with people and larger animals, which results in exceptionally poor image quality when testing mice and other small rodents. Despite this limitation, we have identified seven VFSS parameters that are consistently quantifiable in mice when using a high energy fluoroscope in combination with the new murine VFSS protocol. We recently obtained a low energy fluoroscopy system with exceptionally high imaging resolution and magnification capabilities that was designed for use with mice and other small rodents. Preliminary work using this new system, in combination with the new murine VFSS protocol, has identified 13 swallow parameters that are consistently quantifiable in mice, which is nearly double the number obtained using conventional (i.e., high energy) fluoroscopes. Identification of additional swallow parameters is expected as we optimize the capabilities of this new system. Results thus far demonstrate the utility of using a low energy fluoroscopy system to detect and quantify subtle changes in swallow physiology that may otherwise be overlooked when using high energy fluoroscopes to investigate murine disease models.

This study successfully adapted human videofluoroscopic swallowing study (VFSS) methods for use with murine disease models for the purpose of facilitating translational dysphagia research.

This study adapted human videofluoroscopic swallowing study (VFSS) methods for use with murine disease models for the purpose of facilitating ...

High-frequency Ultrasound Imaging of Mouse Cervical Lymph Nodes

High-frequency ultrasound (HFUS) is widely employed as a non-invasive method for imaging internal anatomic structures in experimental small animal systems. HFUS has the ability to detect structures as small as 30 &#181;m, a property that has been utilized for visualizing superficial lymph nodes in rodents in brightness (B)-mode. Combining power Doppler with B-mode imaging allows for measuring circulatory blood flow within lymph nodes and other organs. While HFUS has been utilized for lymph node imaging in a number of mouse&#160; model systems, a detailed protocol describing HFUS imaging and characterization of the cervical lymph nodes in mice has not been reported. Here, we show that HFUS can be adapted to detect and characterize cervical lymph nodes in mice. Combined B-mode and power Doppler imaging can be used to detect increases in blood flow in immunologically-enlarged cervical nodes. We also describe the use of B-mode imaging to conduct fine needle biopsies of cervical lymph nodes to retrieve lymph tissue for histological&#160; analysis. Finally, software-aided steps are described to calculate changes in lymph node volume and to visualize changes in lymph node morphology following image reconstruction. The ability to visually monitor changes in cervical lymph node biology over time provides a simple and powerful technique for the non-invasive monitoring of cervical lymph node alterations in preclinical mouse models of oral cavity disease.

This protocol describes the application of high-frequency ultrasound (HFUS) for imaging mouse cervical lymph nodes. This technique optimizes visualization and quantification of cervical lymph node morphology, volume and blood flow. Image-guided biopsy of cervical lymph nodes and processing of lymph tissue for histological evaluation is also demonstrated.

High-frequency ultrasound (HFUS) is widely employed as a non-invasive method for imaging internal anatomic structures in experimental small animal ...

A Multicenter MRI Protocol for the Evaluation and Quantification of Deep Vein Thrombosis

We evaluated a magnetic resonance venography (MRV) approach with gadofosveset to quantify total thrombus volume changes as the principal criterion for treatment efficacy&#160;in a multicenter randomized study comparing edoxaban monotherapy with a heparin/warfarin regimen for acute, symptomatic lower extremities deep vein thrombosis (DVT) treatment. We also used a direct thrombus imaging approach (DTHI, without the use of a contrast agent) to quantify fresh thrombus. We then sought to evaluate the reproducibility of the analysis methodology and applicability of using 3D magnetic resonance venography and direct thrombus imaging for the quantification of DVT in a multicenter trial setting. From 10 randomly selected subjects participating in the edoxaban Thrombus Reduction Imaging Study (eTRIS), total thrombus volume in the entire lower extremity deep venous system was quantified bilaterally. Subjects were imaged using 3D-T1W gradient echo sequences before (direct thrombus imaging, DTHI) and 5 min after injection of 0.03 mmol/kg of gadofosveset trisodium (magnetic resonance venography, MRV). The margins of the DVT on corresponding axial, curved multi-planar reformatted images were manually delineated by two observers to obtain volumetric measurements of the venous thrombi. MRV was used to compute total DVT volume, whereas DTHI was used to compute volume of fresh thrombus. Intra-class correlation (ICC) and Bland Altman analysis were performed to compare inter and intra-observer variability of the analysis. The ICC for inter and intra-observer variability was excellent (0.99 and 0.98, p &#60;0.001, respectively) with no bias on Bland-Altman analysis for MRV images. For DTHI images, the results were slightly lower (ICC = 0.88 and 0.95 respectively, p &#60;0.001), with bias for inter-observer results on Bland-Altman plots. This study showed feasibility of thrombus volume estimation in DVT using MRV with gadofosveset trisodium, with good intra- and inter-observer reproducibility in a multicenter setting.

The goal of this study is to use magnetic resonance venography with long-circulating gadolinium-based contrast agent and direct thrombus imaging for quantitative evaluation of DVT volume in a multicenter, clinical trial setting. Inter- and intra-observer variability assessments were conducted, and reproducibility of the protocol was determined.

We evaluated a magnetic resonance venography (MRV) approach with gadofosveset to quantify total thrombus volume changes as the principal criterion for ...

Ferric Chloride-induced Thrombosis Mouse Model on Carotid Artery and Mesentery Vessel

Severe thrombosis and its ischemic consequences such as myocardial infarction, pulmonary embolism and stroke are major worldwide health issues. The ferric chloride injury is now a well-established technique to rapidly and accurately induce the formation of thrombi in exposed veins or artery of small and large diameter. This model has played a key role in the study of the pathophysiology of thrombosis, in the discovery and validation of novel antithrombotic drugs and in the understanding of the mechanism of action of these new agents. Here, the implementation of this technique on a mesenteric vessel and carotid artery in mice is presented. The method describes how to label circulating leukocytes and platelets with a fluorescent dye and to observe, by intravital microscopy on the exposed mesentery, their accumulation at the injured vessel wall which leads to the formation of a thrombus. On the carotid artery, the occlusion caused by the clot formation is measured by monitoring the blood flow with a Doppler probe.

The FeCl3 induced thrombosis model in mice is described herein. A method to monitor thrombus growth by intravital microscopy observation on a mesenteric vessel and by blood flow measurement in the carotid artery is presented.

Severe thrombosis and its ischemic consequences such as myocardial infarction, pulmonary embolism and stroke are major worldwide health issues. The ferric ...