This work was supported by a grant from the Heart and Stroke Foundation of Canada and by The Morris and Bella Fainman Family Foundation. The authors would like to thank Veronique Michaud for her technical help with the VEVO770 ultrasound imaging system.

All procedures were approved by the institutional Animal Care Committee of McGill University Montr&#233;al, QC, Canada. All the equipment required is listed in Table I.
1. Murine (C57BL/6J) IVC Stasis Protocol
<ol>
	<li>Surgery preparation 
	NOTE: This is a survival surgery and therefore, proper aseptic technique needs to be followed at all times. This includes making sure all materials are at hand, cleaning and sterilizing instruments before and between use, and designating a sterile area on the working surface for all sterile material.
	<ol>
		<li>Sterilize all surgical instruments&#160;and materials by autoclave. A glass bead sterilizer is sufficient to sterilize instrument tips between animals, if more than one procedure is being done at one time.</li>
		<li>Anesthetize 8 - 10 week old C57Bl/6 male mice with a mixture of 100% oxygen and 2.5% isoflurane. 
		NOTE: Adjust the flow of oxygen and isoflurane as necessary.</li>
		<li>Confirm anesthetization by pinching the rear paw of the animal between the toes with forceps. No response from the pinch indicates the animal is anesthetized.</li>
		<li>Administer slow release analgesic (buprenorphine) via subcutaneous injection. Give mice a dose of 1 mg of analgesic per 1 kg of body weight (1 mg/kg).</li>
		<li>Place eye ointment on both eyes of the animal to ensure no corneal desiccation for the duration of the procedure.</li>
		<li>Administer 0.2 - 0.5 mL of isotonic fluids per 10 g of body weight subcutaneously.</li>
		<li>Fix the animal in place to the surgery table using surgical tape. Place the animal in a supine position to expose the abdomen. Perform surgery on a heating pad to prevent hypothermia.</li>
		<li>Remove hair from the abdomen of the mouse by application of a hair removal cream for 1 - 2 min. Wipe the abdominal area clean with gauze and distilled water, if necessary.</li>
		<li>Alternatively, remove the hair from the abdomen by shaving with a small electric razor.</li>
		<li>Sterilize the abdomen with&#160;Baxedin (Chlorhexidine Gluconate BP Solution)&#160;&#160;prior to making any incisions. Apply ethanol lightly with gauze to avoid excessive heat loss caused by the evaporation of the alcohol.</li>
	</ol>
	</li>
	<li>Exposure of the Inferior Vena Cava (IVC)
	<ol>
		<li>Using surgical scissors, perform a laparotomy to open the abdominal cavity.
		<ol>
			<li>Lift skin with forceps at the lower abdomen and make a vertical incision parallel to either side of the linea alba. Make the first incision to the left or right of the linea alba, depending on the handedness of the operator. The incision is made away from the midline to assist in ultrasound imaging later on.</li>
			<li>Make a second, horizontal incision at the top of the abdomen. 
			NOTE: Upon penetration into the abdomen, the musculature should be &#8220;tented&#8221; and then penetrated with surgical scissors to prevent any damage to the abdominal organs.</li>
			<li>Repeat the vertical and horizontal incisions on the abdominal muscle layer of the animal.&#160;Be sure to tent the musculature during penetration into the abdomen so as not to damage the organs beneath. 
			Caution: Lift skin and muscle away from intestines and other internal organs when making incisions so as not to puncture them.</li>
		</ol>
		</li>
		<li>Fold the skin and muscle away from the incision to expose the abdominal cavity.</li>
		<li>Apply gauze, moistened with isotonic solution, to the sides of the wound opening and externalize the intestines. Apply pressure to both sides of the abdomen to assist the externalization of the intestines. Use gentle movements with a cotton tip applicator to move the intestines. Soak gauze in isotonic solution and place it over the externalized intestines.&#160;
		<ol>
			<li>Gauze should be pre-moistened prior to placement on tissue.</li>
		</ol>
		</li>
		<li>Move aside any peritoneal fat, and expose the IVC between the renal and iliac veins.</li>
		<li>Make an initial identification of obvious side branches. A side branch will appear as smaller, but significant vein that branches away from the IVC, between the renal and iliac veins. Vasculature can vary greatly between mice. Mice may have side branches present on one or both sides of the IVC.</li>
	</ol>
	</li>
	<li>Ligation of side branches 
	NOTE: If side branches are present, the following procedure is followed for each one. If not present, proceed to step 4: Initial suture placement under the IVC. The ligation site of the IVC will be immediately distal to the left renal vein, regardless of the presence of side branches.
	<ol>
		<li>Perform blunt dissection on each side of the side branch. Blunt dissection is the process of repeatedly opening blunt forceps with gentle pressure, so as to break through fascia without damaging surrounding vessels. 
		Caution: Avoid puncturing or damaging the vein.</li>
		<li>Lift the branch and pass a section of 6-0 silk suture underneath the vein. Always take care when lifting or moving vessels. It is best to grab the fat around the vessels, otherwise there is a risk of damaging the vessel.</li>
		<li>Using suture forceps, make a surgical ligation around the side branch using standard surgical knot tying technique. Full ligation is done to ensure 100% occlusion of the vein. Use at least three tying throws to ensure the ligation is secure.</li>
		<li>Repeat step 1.3 for any remaining side branches.</li>
	</ol>
	</li>
	<li>Dissection of the abdominal aorta from the IVC
	<ol>
		<li>Perform blunt dissection around the IVC immediately distal from the left renal vein. The abdominal aorta is attached to the IVC via fascia, therefore perform initial blunt dissection around both the IVC and abdominal aorta. 
		Caution: Avoid puncturing or damaging the vein. Do not rupture or puncture either vessel. This is the most critical part of the procedure, with the highest risk for vessel damage.</li>
		<li>Continue to apply gentle pressure via blunt dissection until a clear window is made between the aorta and IVC.</li>
	</ol>
	</li>
	<li>Suture placement and IVC ligation
	<ol>
		<li>Pass a section of 6-0 silk suture underneath the IVC and abdominal aorta.</li>
		<li>Locate the suture through the window created between the IVC and aorta. Use forceps to pull the suture and thread it between the abdominal aorta and IVC. Thread the suture through carefully, trying not to pull/puncture the IVC and abdominal aorta.</li>
		<li>Make a surgical ligation around the IVC with suture forceps, ensuring a total occlusion of the vein. Again, make the ligation immediately distal from the left renal vein.</li>
		<li>Make three suture throws to secure the ligation. 
		NOTE: It is also possible to pass the suture underneath the IVC and aorta prior to separating the two as an alternative option.&#160;Alternatively, 7-0 Prolene sutures can be used for ligation of the side branches and the IVC.</li>
	</ol>
	</li>
	<li>Wound closure and post-operative care
	<ol>
		<li>Verify that the abdominal aorta has been left uninterrupted and that all side branches have been ligated. The IVC will appear dilated distal from the ligation site and no blood flow will be visible.</li>
		<li>Place all peritoneal fat and the intestines back in the abdominal cavity.</li>
		<li>Close the wound using suture forceps and 6-0 silk suture. Use a running suture (simple continuous) and ensure that the suture will not be too tight. A tight suture will rupture when the animal wakes up and moves around its cage.
		<ol>
			<li>Alternatively, use PDS, Ethilon, Vicryl, Dexon, or stainless steel clips for wound closure.</li>
		</ol>
		</li>
		<li>Suture the layer of abdominal muscle first using appropriate suture tying technique.</li>
		<li>Repeat suture technique for the skin and verify wound closure is sufficient. 
		NOTE: If using the same suture needle and thread for multiple animals, sterilize both in 70% ethanol prior to each use.</li>
		<li>Remove the animal from anesthetic gas and place them in a 34 &#176;C incubator for at least 30 minutes.</li>
		<li>Again, administer 0.2 - 0.5 mL of isotonic fluids per 10 g of body weight subcutaneously. Fluids can be given in following days to maintain pre-operative body weight, if necessary.</li>
		<li>Monitor the animal until recovery to ensure successful surgery and assess the overall health of the animal. 
		Caution: Expect a slight hunched posture for an invasive procedure such as this, but the animal will behave normally as early as 30 minutes post-operation.</li>
		<li>Place the animal in its own cage and do not place it with other animals until fully recovered.</li>
		<li>Re-administer analgesic daily for up to 48 h&#160;postop.</li>
		<li>Place food on the floor of the cage for the animal while in recovery. Other special care is generally not needed.</li>
		<li>Examine the wound site for redness, swelling, or discharge and any other signs of infection.</li>
		<li>If necessary, remove sutures after 7 - 10 days.</li>
		<li>Perform euthanasia immediately if surgery is unsuccessful (e.g., significant vessel damage and/or blood loss) or animal does not improve with post-operative care.&#160;Euthanasia is generally performed at 48 h&#160;postop. For longer experiments, use 7-0 Prolene to ligate side branches and the IVC and 5-0 Vicryl sutures to close the abdominal cavity.
		<ol>
			<li>Perform euthanasia by anesthetization of the animal in an induction as previously described, except at 5% isoflurane. This is followed by asphyxiation with 100% CO2 while anesthetized.</li>
			<li>After confirmation of euthanasia by asphyxiation (loss of heartbeat and no breathing), perform cervical dislocation to ensure complete euthanasia.</li>
		</ol>
		</li>
	</ol>
	</li>
</ol>
2. High Frequency Ultrasound Protocol
NOTE: This protocol is adapted from the Lady Davis Institute Rodent Phenotyping Core SOP for high frequency ultrasound imaging. This protocol is carried out 24 h&#160;post-operative, but can be done sooner as long as the animal is responding well to the surgery. The protocol can be carried out at any time on healthy mice and is often done so to compare before and after surgery.
<ol>
	<li>Preparation of Animal
	<ol>
		<li>Place the animal in an induction chamber and anesthetize the animal with a mixture of 100% oxygen and 2.5% isoflurane.</li>
		<li>Turn on heart rate and temperature monitor. Position heat lamp above table to keep animal warm during procedure.</li>
		<li>Remove the animal from the induction chamber and apply eye lubricant to prevent corneal desiccation. Position the animal on the analysis platform and affix the anesthetic tube to the animal&#39;s mouth/nose.</li>
		<li>Warm ultrasound gel to 37 &#176;C and place the gel on the abdomen of the animal. The ultrasound gel is warmed using a water-heating system that pumps warm water through coils that are wrapped around the bottle of gel.</li>
		<li>For heart rate measurements (if needed), put electrode gel on the 4 electrodes of the analysis platform and fix the paws of the animal to the platform/electrodes with surgical tape. Place the animal in a supine position for imaging of the IVC. The ultrasound imaging system includes an analysis platform that is warmed and contains electrodes for monitoring the animal.</li>
		<li>For temperature measurements, put electrode gel on thermometer and insert into the rectum of the animal.</li>
		<li>Adjust heat lamp and isoflurane as necessary to ensure that animal is comfortable, has a respiration between 30 - 70 bpm, and is kept at 37 &#176;C.</li>
		<li>If necessary, shave abdomen of animal or remove hair with hair removal cream. Apply for 1 - 2 minutes and wipe clean with gauze and distilled water. 
		NOTE: Too much hair can affect the quality of the images.</li>
		<li>Place gauze under or to the side of the animal to catch any excess ultrasound gel.</li>
	</ol>
	</li>
	<li>Imaging the IVC and Thrombus
	<ol>
		<li>Turn on the imaging software and begin a new study.</li>
		<li>Select an appropriate scanhead for imaging. The scanhead used here is for abdominal organs (Frequency: 35 MHz, Focal length: 12.8 mm).</li>
		<li>Record any relevant information about the animal. This may include animal ID, sex, weight, birthdate, strain, etc.</li>
		<li>Lower the scanhead down until it is touching the ultrasound gel on the animal.</li>
		<li>Start the scanhead probe to begin 2-D imaging of the animal. In this mode, images are presented as two dimensional, in varying shades of gray.</li>
		<li>Locate the IVC and abdominal aorta by moving the mouse platform throughout the x- and/or y-plane, while also moving the scanhead through the z-plane. 
		NOTE: Blood vessels will appear with their walls white and the blood inside nearly black. The abdominal aorta will pulsate intensely and have thicker walls, while the IVC will have thinner walls and compress/expand easily on the image screen when the scanhead is moved up and down. The ligation site will be apparent, as the vein will be dilated and the vein wall will come to a point. Any thrombus formed inside will be solid under ultrasound imaging and will not compress easily like the vein wall does. The vein will compress only above the ligation, where no thrombus is formed.</li>
		<li>Upon locating the IVC and the ligation site, center them on the focal point of the ultrasound, in order to achieve the most accurate image.
		<ol>
			<li>If the image is too dark to see clearly, adjust the gel and remove any air bubbles with a cotton tip applicator.</li>
			<li>If necessary, tilt the scanhead 45&#176; to the left or right and readjust the animal platform to see the vein clearly. This is done if there is interference with the scanhead. In this scenario, the vein needs to be imaged at a side-angle to avoid the suture. 
			NOTE: Image interference is most often caused by dark patches of skin on the animal, poor wound suture placement, or bubbles in the ultrasound gel.</li>
		</ol>
		</li>
		<li>Using the software, take photos of any desired structures and make measurements using any measurement tools. Common measurements include cross-sectional area of the thrombus or width of the ligation site.</li>
		<li>Change the imaging mode to Pulse-wave Doppler to make blood flow measurements. This mode allows the imaging of vascular blood flow and the measurement of flow velocity, among other variables.</li>
		<li>Tilt the analysis platform to lower the posterior end of the animal.</li>
		<li>Change the angle/position of the scanhead so that it will contact the gel at an angle of about 30&#176; from the posterior end of the animal.</li>
		<li>Reposition the animal platform and scanhead. Relocate the IVC and ligation if necessary.</li>
		<li>Make blood flow measurements around the ligation site to confirm stasis and take any images desired. Common measurements of blood flow include average velocity, peak velocity, or median velocity.</li>
		<li>Save any measurements or images taken.</li>
	</ol>
	</li>
	<li>Clean-up and animal recovery
	<ol>
		<li>Raise the scanhead to initial position and reposition animal platform to its initial position, as well.</li>
		<li>Remove excess gel off the animal with gauze. Do this gently so that the wound suture will not be ruptured.</li>
		<li>Remove the thermometer from the animal&#39;s rectum and take the animal off the platform.</li>
		<li>Place the animal in its cage and a 34 &#176;C incubator.</li>
		<li>Monitor the animal until recovery. The procedure is very minor and non-invasive. The animal will wake up quickly. Continue monitoring animal in the following days in accordance with section 1.6 of this protocol.</li>
		<li>Clean the animal platform with gauze and disinfectant.</li>
		<li>Wipe the scanhead clean with gauze and distilled water. 
		Caution: Clean the scanhead very gently and only with distilled water.</li>
		<li>Repeat step 2 for the next animal.
		<ol>
			<li>If done with all subjects, verify that all images and recordings have been saved. Then turn off all equipment and software.</li>
		</ol>
		</li>
	</ol>
	</li>
</ol>

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The authors have nothing to disclose.

There are several critical steps for successful venous thrombus formation using the stasis model. Induction of vein thrombosis is more challenging in old mice due to the accumulation of fat surrounding the inferior vena cava and the aorta. Ideally, mice undergoing this procedure should be 8 - 10-weeks-old. Great care should be taken not to induce endothelial damage in the IVC during the blunt dissection and ligature. In addition, it is crucial to keep the animal in a 34 &#176;C incubator for at least 30 minutes after the...

Venous thromboembolism (VTE), which is comprised of deep venous thrombosis (DVT) and pulmonary embolism, is the third leading cause of cardiovascular death after myocardial infarction and stroke. It is a common condition affecting 1 - 2% of the population, with an annual incidence of 1 in 5001. VTE can lead to: 1) death through pulmonary embolism; 2) post-thrombotic syndrome, characterized by chronic leg pain, swelling, and ulceration; or 3) chronic pulmonary hypertension resulting in significant chronic respiratory compromise. VTE is a multi-factorial disease and may result from stasis of blood flow, damage to vessel walls, and/or hypercoagulable states due to a disruption of the balance between the coagulation and the fibrinolytic systems, as it has been described over a hundred years ago by Virchow and is known as the Virchow&#39;s Triad.
Because in most cases it is impossible to obtain human DVT samples, researchers have developed experimental animal models of DVT. Several animals including rat2, mouse3, rabbit4, pig5, dog6, and non-human primate7 have been used. Mice can be genetically modified and are the most frequently used animal to study DVT. However, as in all animals, spontaneous DVT is not observed in mice. Thus, physical or chemical alterations of the vein wall are used to create thrombosis in mice. We have previously used the ferric chloride model to induce thrombosis in the inferior vena cava (IVC) of mice8,9,10. This model has the advantage of reliably producing occlusive thrombi within minutes and can be used to investigate the role of anti-coagulant and anti-platelet drugs during acute DVT. However, it is a terminal procedure. Thus, to study acute and chronic DVT, the stasis model is more suitable. In this model, thrombus formation is induced by the complete interruption of blood flow in the IVC, one of the factors in Virchow&#39;s triad for DVT development. This model can be used to study DVT formation and resolution, which is an advantage compared to the FeCl3 model11.
We have developed a non-invasive method to follow thrombus formation and resolution over time using a micro-imaging high-frequency ultrasound system12. We have previously demonstrated that measurement of venous thrombosis by ultrasound correlates favorably with thrombi obtained pathologically. We have confirmed in two subsequent studies that measurements obtained with ultrasound correlate with thrombus weight and thrombus area quantified by histochemistry9,10. More importantly, we have showed that high frequency ultrasound can be used to monitor the formation of deep venous thrombosis in mice12. It may also be used to quantify thrombus resolution in a non-invasive way.
Here, we will describe the protocol allowing thrombus formation using the stasis-induced thrombosis mouse model and how thrombus formation can be monitored non-invasively over time using high frequency ultrasound.

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deep vein thrombosis induced by stasis in mice monitored by high frequency ultrasonography

Venous thrombosis is a common condition affecting 1 - 2% of the population, with an annual incidence of 1 in 500. Venous thrombosis can lead to death through pulmonary embolism or results in the post-thrombotic syndrome, characterized by chronic leg pain, swelling, and ulceration, or in chronic pulmonary hypertension resulting in significant chronic respiratory compromise. This is the most common cardiovascular disease after myocardial infarction and ischemic stroke and is a clinical challenge for all medical disciplines, as it can complicate the course of other disorders such as cancer, systemic disease, surgery, and major trauma.
Experimental models are necessary to study these mechanisms. The stasis model induces consistent thrombus size and a quantifiable amount of thrombus. However, it is necessary to systematically ligate side branches of the inferior vena cava to avoid variability in thrombus sizes and any erroneous data interpretation. We have developed a non-invasive technique to measure thrombus size using ultrasonography. Using this technique, we can assess thrombus development and resolution over time in the same animal. This approach limits the number of mice required for quantification of venous thrombosis consistent with the principle of replacement, reduction, and refinement of animals in research. We have demonstrated that thrombus weight and histological analysis of thrombus size correlate with measurement obtained with ultrasonography. Therefore, the current study describes how to induce deep vein thrombosis in mice using the inferior vena cava stasis model and how to monitor it using high frequency ultrasound.

Stasis venous thrombosis model
In the stasis model, mice are anesthetized, and an incision is made to expose the inferior vena cava (IVC). The incision is made on the left or right side of the mouse instead of a midline laparotomy in a way that would not interfere with the ultrasound probe. The abdominal muscles and the skin are fold back to expose the IVC (Figure 1). First, side br...

Watch this Scientific Journal Video about Deep Vein Thrombosis Induced by Stasis in Mice Monitored by High Frequency Ultrasonography at JoVE.com

Deep Vein Thrombosis Induced by Stasis in Mice Monitored by High Frequency Ultrasonography

The present protocol describes the steps to obtain venous thrombosis using a stasis model. In addition, we are using a non-invasive method to measure thrombus formation and resolution over time.

Venous thrombosis is a common condition affecting 1 - 2% of the population, with an annual incidence of 1 in 500. Venous thrombosis can lead to death ...

deep-vein-thrombosis-induced-stasis-mice-monitored-high-frequency

Universidad de Cartagena UDEC

Research

JoVE Journal

Immunology and Infection

9.8K Views.  McGill University. The present protocol describes the steps to obtain venous thrombosis using a stasis model. In addition, we are using a non-invasive method to measure thrombus formation and resolution over time.

Video: Deep Vein Thrombosis Induced by Stasis in Mice Monitored by High Frequency Ultrasonography

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 ...

Medicine

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 ...

Optimized Venous Thrombosis Assay in a Mouse Cancer Model Using Vascular Clips

Venous Thrombosis Assay in a Mouse Model of Cancer

This methodology paper highlights the surgical nuances of a rodent model of venous thrombosis, specifically in the context of cancer-associated thrombosis (CAT). Deep venous thrombosis is a common complication in cancer survivors and can be potentially fatal. The current murine venous thrombosis models typically involve a complete or partial mechanical occlusion of the inferior vena cava (IVC) using a suture. This procedure induces a total or partial stasis of blood and endothelial damage, triggering thrombogenesis. The current models have limitations such as higher variability in clot weights, significant mortality rate, and prolonged learning curve. This report introduces surgical refinements using vascular clips to address some of these limitations. Using a syngeneic colon cancer xenograft mouse model, we employed customized vascular clips to ligate the infrarenal vena cava. These clips allow residual lip space similar to a 5-0 polypropylene suture after IVC ligations. Mice with the suture method served as controls. The vascular clip method resulted in a consistent reproducible partial vascular occlusion and greater clot weights with less variability than the suture method. The larger clot weights, greater clot mass, and clot to the IVC luminal surface area were expected due to the higher pressure profile of the vascular clips compared to a 6-0 polypropylene suture. The approach was validated by gray scale ultrasonography, which revealed consistently greater clot mass in the infrarenal vena cava with vascular clips compared to the suture method. These observations were further substantiated with the immunofluorescence staining. This study offers an improved method to generate a venous thrombosis model in mice, which can be employed to deepen the mechanistic understanding of CAT and in translational research such as drug discovery.

Author Spotlight: Advancing Cancer Associated Thrombosis Research in Rodent Models 

This article aims to present an optimized method for assessing venous thrombosis in a mouse cancer model, using vascular clips to achieve venous ligation. Optimization minimizes variability in thrombosis-related measurements and enhances relevance to human cancer-associated venous thrombosis.

This methodology paper highlights the surgical nuances of a rodent model of venous thrombosis, specifically in the context of cancer-associated thrombosis ...

Cancer Research

Stenosis of the Inferior Vena Cava: A Murine Model of Deep Vein Thrombosis

Deep vein thrombosis (DVT) and its devastating complication, pulmonary embolism, are a severe health problem with high mortality. Mechanisms of thrombus formation in veins remain obscure. Lack of mobility (e.g., after surgery or long-haul flights) is one of the main factors leading to DVT. The pathophysiological consequence of the lack of mobility is blood flow stagnation in venous valves. Here, a model is described that mimics such flow disturbance as a thrombosis-driving factor. In this model, partial flow restriction (stenosis) in the inferior vena cava (IVC) is created. Closure of about 90% of the IVC lumen for 48 h results in development of thrombi structurally similar to those in humans. The similarities are: i) most of the thrombus volume is red, i.e., consists of red blood cells and fibrin, ii) presence of a white part (lines of Zahn), iii) non-denuded endothelial monolayer, iv) elevated plasma D-Dimer levels, and v) possibility to prevent thrombosis by low molecular weight heparin. Limitations include variable size of thrombi and the fact that a certain percentage of wild-type mice (0 - 35%) may not produce a thrombus. In addition to visual observation and measurement, thrombi may be visualized by non-invasive technologies, such as ultrasonography, which allows for monitoring the dynamics of thrombus development. At shorter time points (1 - 6 h), intravital microscopy may be applied to directly observe events (e.g., recruitment of cells to the vessel wall) preceding thrombus formation. Use of this method by several teams around the world has made it possible to uncover basic mechanisms of DVT initiation and identify potential targets that might be beneficial for its prevention.

We describe here stenosis in the inferior vena cava as a murine model of deep vein thrombosis. This model recapitulates blood flow restriction, one of the major triggers of venous thrombosis in humans.

Deep vein thrombosis (DVT) and its devastating complication, pulmonary embolism, are a severe health problem with high mortality. Mechanisms of thrombus ...

Methods for Acute and Subacute Murine Hindlimb Ischemia

Peripheral artery disease (PAD) is a leading cause of cardiovascular morbidity and mortality in developed countries, and animal models that reliably reproduce the human disease are necessary to develop new therapies for this disease. The mouse hindlimb ischemia model has been widely used for this purpose, but the standard practice of inducing acute limb ischemia by ligation of the femoral artery can result in substantial tissue necrosis, compromising investigators&#39; ability to study the vascular and skeletal muscle tissue responses to ischemia. An alternative approach to femoral artery ligation is the induction of gradual femoral artery occlusion through the use of ameroid constrictors. When placed around the femoral artery in the same or different locations as the sites of femoral artery ligation, these devices occlude the artery over 1 - 3 days, resulting in more gradual, subacute ischemia. This results in less substantial skeletal muscle tissue necrosis, which may more closely mimic the responses seen in human PAD. Because genetic background influences outcomes in both the acute and subacute ischemia models, consideration of the mouse strain being studied is important in choosing the best model. This paper describes the proper procedure and anatomical placement of ligatures or ameroid constrictors on the mouse femoral artery to induce subacute or acute hindlimb ischemia in the mouse.

Surgical induction of hindlimb ischemia in the mouse is useful to examine angiogenesis, however this is compromised in certain inbred mouse strains that display marked ischemia-induced tissue necrosis. Methods are described to induce subacute limb ischemia using ameroid constrictors to circumvent this problem through the induction of gradual arterial occlusion.

Peripheral artery disease (PAD) is a leading cause of cardiovascular morbidity and mortality in developed countries, and animal models that reliably ...

Intravital Microscopy and Thrombus Induction in the Earlobe of a Hairless Mouse

Thrombotic complications of vascular diseases are one leading cause of morbidity and mortality in industrial nations. Due to the complex interactions between cellular and non-cellular blood components during thrombus formation, reliable studies of the physiology and pathophysiology of thrombosis can only be performed in vivo. Therefore, this article presents an ear model in hairless mice and focuses on the in vivo analysis of microcirculation, thrombus formation, and thrombus evolution. By using intravital fluorescence microscopy and the intravenous (iv) application of the respective fluorescent dyes, a repetitive analysis of microcirculation in the auricle can easily be performed, without the need for surgical preparation. Furthermore, this model can be adapted for in vivo studies of different issues, including wound healing, reperfusion injury, or angiogenesis. In summary, the ear of hairless mice is an ideal model for the in vivo study of skin microcirculation in physiological or pathophysiological conditions and for the evaluation of its reaction to different systemic or topical treatments.

The ear model of the hairless SKH1-Hrhr mouse enables intravital fluorescence microscopy of microcirculation and phototoxic thrombus induction without prior surgical preparation in the examined microvascular bed. Therefore, the ear of the hairless mouse is an excellent in vivo model to study the complex interactions during microvascular thrombus formation, thrombus evolution, and thrombolysis.

Thrombotic complications of vascular diseases are one leading cause of morbidity and mortality in industrial nations. Due to the complex interactions ...

Murine Model of Central Venous Stenosis using Aortocaval Fistula with an Outflow Stenosis

Central venous stenosis is an important entity contributing to arteriovenous fistula (AVF) failure. A murine AVF model was modified to create a partial ligation of the inferior vena cava (IVC) in the outflow of the fistula, mimicking central venous stenosis. Technical aspects of this model are introduced. The aorta and IVC are exposed, following an abdominal incision. The infra-renal aorta and IVC are dissected for proximal clamping, and the distal aorta is exposed for puncture. The IVC at the midpoint between the left renal vein and the aortic bifurcation is carefully dissected to place an 8-0 suture beneath the IVC. After clamping the aorta and IVC, an AVF is created by puncturing the infra-renal aorta through both walls into the IVC with a 25 G needle, followed by ligating a 22 G intra-venous (IV) catheter and IVC together. The catheter is then removed, creating a reproducible venous stenosis without occlusion. The aorta and IVC are unclamped after confirming primary hemostasis. This novel model of central vein stenosis is easy to perform, reproducible, and will facilitate studies on AVF failure.

An aortocaval fistula was created by puncturing the murine infra-renal aorta through both walls into the inferior vena cava and was followed by creation of a stenosis in its outflow via partial ligation of the inferior vena cava. This reproducible model can be used to study central venous stenosis.

Central venous stenosis is an important entity contributing to arteriovenous fistula (AVF) failure. A murine AVF model was modified to create a partial ...

Analysis of Cerebral Vasospasm in a Murine Model of Subarachnoid Hemorrhage with High Frequency Transcranial Duplex Ultrasound

Cerebral vasospasm that occurs in the weeks after subarachnoid hemorrhage, a type of hemorrhagic stroke, contributes to delayed cerebral ischemia. A problem encountered in experimental studies using murine models of SAH is that methods for in vivo monitoring of cerebral vasospasm in mice are lacking. Here, we demonstrate the application of high frequency ultrasound to perform transcranial Duplex sonography examinations on mice. Using the method, the internal carotid arteries (ICA) could be identified. The blood flow velocities in the intracranial ICAs were accelerated significantly after induction of SAH, while blood flow velocities in the extracranial ICAs remained low, indicating cerebral vasospasm. In conclusion, the method demonstrated here allows functional, noninvasive in vivo monitoring of cerebral vasospasm in a murine SAH model.

The aim of this manuscript is to present a sonography-based method that allows in vivo imaging of blood flow in cerebral arteries in mice. We demonstrate its application to determine changes in blood flow velocities associated with vasospasm in murine models of subarachnoid hemorrhage (SAH).

Cerebral vasospasm that occurs in the weeks after subarachnoid hemorrhage, a type of hemorrhagic stroke, contributes to delayed cerebral ischemia. A ...

Neuroscience

High-Resolution Three-Dimensional Imaging of the Footpad Vasculature in a Murine Hindlimb Gangrene Model

Peripheral arterial disease (PAD) is a significant cause of morbidity resulting from chronic exposure to atherosclerotic risk factors. Patients suffering from its most severe form, chronic limb-threatening ischemia (CLTI), face substantial impairments to daily living, including chronic pain, limited walking distance without pain, and nonhealing wounds. Preclinical models have been developed in various animals to study PAD, but mouse hindlimb ischemia remains the most widely used. There can be significant variation in response to ischemic insult in these models depending on the mouse strain used and the site, number, and means of arterial disruption. This protocol describes a unique method combining femoral artery and vein electrocoagulation with the administration of a nitric oxide synthase (NOS) inhibitor to reliably induce footpad gangrene in Friend Virus B (FVB) mice that resembles the tissue loss of CLTI. While traditional means of assessing reperfusion such as laser Doppler perfusion imaging (LDPI) are still recommended, intracardiac perfusion of the lipophilic dye 1,1'-dioctadecyl-3,3,3',3'-tetramethylindocarbocyanine perchlorate (DiI) is used to label the vasculature. Subsequent whole-mount confocal laser scanning microscopy allows for high-resolution, three-dimensional (3D) reconstruction of footpad vascular networks that complements traditional means of assessing reperfusion in hindlimb ischemia models.

The present protocol describes a unique, clinically relevant model of peripheral arterial disease that combines femoral artery and vein electrocoagulation with the administration of a nitric oxide synthase inhibitor to induce hindlimb gangrene in FVB mice. Intracardiac DiI perfusion is then used for high-resolution, three-dimensional imaging of the footpad vasculature.

Peripheral arterial disease (PAD) is a significant cause of morbidity resulting from chronic exposure to atherosclerotic risk factors. Patients suffering ...

Intravital Fluorescence Microscopy to Study Microvascular Thrombus Formation

Source: Str&#252;der, D., et al. <a href="https://appjove.unicartagenaproxy.elogim.com/v/55174">Intravital Microscopy and Thrombus Induction in the Earlobe of a Hairless Mouse.</a> J. Vis. Exp. (2017)
This video demonstrates the intravital fluorescence microscopy technique to study thrombus formation in a mouse model. The mouse is injected with a fluorophore-labeled polysaccharide to monitor blood circulation. Upon illumination with a high-intensity light, the phototoxic induction of an injury in the microvasculature of the ear leads to the formation of a thrombus, which is confirmed by visualizing the obstruction in the blood flow.