This work was supported by US National Institute of Health (NIH) Grant R01-HL128406; the United States Department of Veterans Affairs Biomedical Laboratory Research and Development Program Merit Review Award I01-BX002336; as well as with the resources and the use of facilities at the VA Connecticut Healthcare System, West Haven, CT.

All experiments were performed with approval from the Yale University Institutional Animal Care and Use Committee (IACUC).
1. Anesthesia and pre-operative procedures
<ol>
	<li>Sterilize all surgical instruments and materials by autoclaving. Turn on the thermal support device to be certain it is warm (40&#8211;42 &#176;C).</li>
	<li>Place a 9&#8211;11-week old C57BL/6 mouse into an acrylic induction chamber and anesthetize it with vaporized 2.5% isoflurane and 0.8 L/min oxygen. Anesthesia induction takes about 3 min.</li>
	<li>Remove the mouse from the chamber. Confirm a deep plane of anesthesia by a toe pinch, an ear pinch and a tail pinch. Place the mouse in supine position on the surgical area and deliver 2.5% isoflurane using silicone mask. Provide buprenorphine at 0.1 mg/kg analgesic and apply ophthalmic ointment to the eyes.</li>
	<li>Remove fur from the ventral side of the neck to the lower abdomen using Nair, a hair remover.</li>
	<li>Cleanse and disinfect the surgical site by using a two-step scrub with 10% povidone-iodine and 70% isopropanol. Apply a surgical drape.</li>
</ol>
2. Operative procedures
<ol>
	<li>Exposure of the clamp and puncture sites
	<ol>
		<li>Prepare sterile instruments and wear sterile gloves to maintain sterility throughout the surgery.</li>
		<li>Make a skin-deep midline abdominal incision with a scalpel from the level of the lower liver edge to just above the pubis. Cut through the musculature with scissors to open the abdominal cavity.</li>
		<li>Insert a retractor into the abdomen and pull out the bowels to the right side. Keep them moist by wrapping in a saline-soaked gauze. Retreive the bladder and the seminal vesicles (in male mice) and pull them out to the caudal side. Dissect the mesentery between the rectum and retroperitoneum with a micro-needle holder to obtain a full view of the aorta and IVC.</li>
		<li>Dissect the infra-renal aorta and IVC en bloc from the lateral and dorsal surrounding retroperitoneal tissues with a micro-needle holder to cross-clamp them together.</li>
		<li>Dissect the surrounding tissues to expose the aortic puncture site at approximately three-quarters of the distance from the left renal vein to the aortic bifurcation.</li>
	</ol>
	</li>
	<li>IVC dissection
	<ol>
		<li>Dissect between the infra-renal IVC and aorta immediately distal to the left renal vein. Extend the dissection distally to the halfway between the left renal vein and the aortic bifurcation, so that the infra-renal IVC, both upstream and downstream to the stenosis, can be observed postoperatively. 
		NOTE: Blunt dissection between the IVC and aorta should be performed from immediately distal to the left renal vein where the connective tissue between the IVC and the aorta is relatively loose.</li>
		<li>Make a window to separate the IVC from the aorta at that level and dissect the IVC from the surrounding tissue. Place an 8-0 polyamide monofilament suture firstly beneath the IVC and aorta (Figure 1A), then position the suture beneath only the IVC (Figure 1B) by pulling the suture end through the window. 
		NOTE: Since the IVC is fragile, dissecting along the aortic adventitia is useful to make a window to prevent the IVC as well as small IVC or aortic branches from being damaged. If bleeding occurs, it is likely to be uncontrollable. If the IVC has distinct side branches, place an 8-0 suture distally to the branches.</li>
	</ol>
	</li>
	<li>AVF creation
	<ol>
		<li>Bend a 25 G needle to a 45&#8211;60&#176; angle at a point ~4 mm from the needle tip.</li>
		<li>Clamp the infra-renal aorta and IVC by applying a microsurgical clip.</li>
		<li>Rotate the aorta medially and caudally by grasping the connective tissue surrounding the bifurcation to expose the puncture site of the aorta stretched it slightly to the ventral side.</li>
		<li>Holding the aorta in a suitable position, puncture through the aorta into the IVC using the prepared 25 G needle (Figure 1C).</li>
		<li>Release the aorta and cover the puncture site with the surrounding tissue pulling up from the left side of the aorta. Take out the needle and press the puncture site gently using a cotton-tipped swab for hemostasis.</li>
	</ol>
	</li>
	<li>Creation of the IVC stenosis 
	<ol>
		<li>Place a tip of a 22 G IV catheter (see the Table of Materials) onto the IVC longitudinally. Ligate the IV catheter and IVC together with an 8-0 suture (Figure 1D), and then remove the IV catheter.</li>
		<li>Confirm primary hemostasis (Figure 1E) and then unclamp the aorta and IVC. Cover the puncture site 1 min more to ensure hemostasis. 
		NOTE: Do not clamp for too long so as to avoid IVC thrombosis distal to the stenosis.</li>
		<li>Return organs to their original positions and close the abdomen with 6-0 sutures.</li>
	</ol>
	</li>
</ol>
3. Post-operative procedures
<ol>
	<li>After closure of the abdominal wound, discontinue isoflurane inhalation. Put the mouse into an individual bedding-free cage and place the cage on a thermal support device to prevent hypothermia. 
	NOTE: The mouse is observed until they achieve and maintain sternal recumbency. Apply post-operative care including analgesia and wound care in accordance with recommendations of the local IACUC. For analgesia we use buprenorphine at 0.1 mg/kg intrasmuscularly every 12 h for 48&#160;h following the surgical procedures and thereafter as needed.</li>
	<li>Confirm the AVF patency postoperatively by using Doppler ultrasound (see the Table of Materials). In addition, measure other vessel and flow characteristics as required.</li>
</ol>

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D., 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=Eph-B4+regulates+adaptive+venous+remodeling+to+improve+arteriovenous+fistula+patency.">Eph-B4 regulates adaptive venous remodeling to improve arteriovenous fistula patency.</a> Scientific Reports. 7 (1), 15386 (2017).</li><li>Payne, H., Brill, A. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Stenosis+of+the+Inferior+Vena+Cava:+A+Murine+Model+of+Deep+Vein+Thrombosis.">Stenosis of the Inferior Vena Cava: A Murine Model of Deep Vein Thrombosis.</a> J Vis Exp. (130), (2017).</li><li>Nam, D., 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=Partial+carotid+ligation+is+a+model+of+acutely+induced+disturbed+flow,+leading+to+rapid+endothelial+dysfunction+and+atherosclerosis.">Partial carotid ligation is a model of acutely induced disturbed flow, leading to rapid endothelial dysfunction and atherosclerosis.</a> American Journal of Physiology: Heart and Circulatory Physiology. 297 (4), H1535-H1543 (2009).</li><li>Ene-Iordache, B., Remuzzi, A. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Disturbed+flow+in+radial-cephalic+arteriovenous+fistulae+for+haemodialysis:+low+and+oscillating+shear+stress+locates+the+sites+of+stenosis.">Disturbed flow in radial-cephalic arteriovenous fistulae for haemodialysis: low and oscillating shear stress locates the sites of stenosis.</a> Nephrology, Dialysis, Transplantation. 27 (1), 358-368 (2012).</li><li>Yamamoto, K., 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=Disturbed+shear+stress+reduces+Klf2+expression+in+arterial-venous+fistulae+in+vivo.">Disturbed shear stress reduces Klf2 expression in arterial-venous fistulae in vivo.</a> Physiological reports. 3, (2015).</li><li>Remuzzi, A., Ene-Iordache, B. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Novel+paradigms+for+dialysis+vascular+access:+upstream+hemodynamics+and+vascular+remodeling+in+dialysis+access+stenosis.">Novel paradigms for dialysis vascular access: upstream hemodynamics and vascular remodeling in dialysis access stenosis.</a> Clinical Journal of the American Society of Nephrology. 8 (12), 2186-2193 (2013).</li></ol>

The authors have nothing to disclose.

The murine AVF model has been used to study the basic mechanisms and molecular events leading to AVF maturation13,14. In this study, we modified an established murine AVF model to create a novel murine aortocaval fistula model with an IVC stenosis in the outflow tract of the fistula. Our ligation model is similar to several previously described murine models that use vascular ligation. A murine model of deep vein thrombosis was created using partial IVC ligation ...

Arteriovenous fistulae (AVF) are the most common accesses for hemodialysis, with superior patency and reduced infection compared to other accesses such as grafts or central venous catheters. However, up to 60% of AVF fail to mature1,2,3; a recent systematic review reported that primary patency rates at 1 year were only 60%4. Stenosis along the venous outflow predominantly causes failure of AVF maturation5,6. There are certain characteristic locations prone to stenosis proximal to the fistula: the juxtaanastomotic swing segment for the radiocephalic fistula, the cephalic arch region for the brachiocephalic fistula and the central vein for the fistula with previously placed ipsilateral subclavian or internal jugular vein catheters7,8.
Central venous stenosis is often asymptomatic in patients without an AVF, but can cause ipsilateral extremity edema by venous hypertension as well as failure of fistula maturation when challenged by fistula flow9. The pathophysiology of central venous stenosis is most likely related to inflammation and the activated coagulation cascade after device placement. Furthermore, constant movement of the catheter tip as well as increased flow from the fistula can alter shear stress, resulting in platelet deposition and venous wall thickening10. To understand the basic mechanisms underlying AVF failure caused by central venous stenosis, an animal model mimicking central venous stenosis with an AVF is required.
We have established a murine aortocaval fistula model that is easy to perform and master and recapitulates the clinical course of human AVF.11 We applied the concepts and technique of several previously established murine models to create a novel murine AVF model with venous stenosis. We introduce a murine aortocaval fistula model with an IVC stenosis in the outflow fistula that can be used for the study of central venous stenosis.

<table><tbody><tr><td>20-60 Mhz scan head</td><td>VisualSonics Inc.</td><td>RMV-704</td><td></td></tr><tr><td>8-0 Sterile Micro Suture, 6mm (140 &amp;micro;), 3/8 Circle, TAP Point Needle</td><td>AROSuture</td><td>T06A08N14-13</td><td>polyamide monofilament sutures</td></tr><tr><td>Induction Chamber, 2 Liter&lt;br/&gt; 3.75&quot;W x 9.00&quot;D x 3.75&quot;H</td><td>VetEquip</td><td>941444</td><td></td></tr><tr><td>Isoflo, Isoflurane liquid</td><td>Zoetis</td><td>26675-46-7</td><td></td></tr><tr><td>Mice, C57BL/6J</td><td>The Jackson Laboratory</td><td>664</td><td></td></tr><tr><td>Pet Bed Microwave Heating Pad</td><td>Snuggle Safe</td><td>6250</td><td></td></tr><tr><td>PrecisionGlide Needle 25G</td><td>BD</td><td>305122</td><td></td></tr><tr><td>Surflo I.V. Catheter 22G</td><td>Terumo</td><td>SR-OX2225CA</td><td>0.85mm outer diameter</td></tr><tr><td>Vascular clamp</td><td>Roboz Surgical Instrument</td><td>RS-5424</td><td></td></tr><tr><td>Vevo770 High Resolution Imaging System</td><td>VisualSonics Inc.</td><td>770</td><td></td></tr></tbody></table>

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.

Male mice underwent the operation mentioned above to create both an AVF and an IVC stenosis. Control mice underwent only laparotomy and dissection of the tissues surrounding the IVC, e.g., a sham procedure, or only creation of an IVC stenosis without simultaneous creation of an AVF.
The IVC was observed with Doppler ultrasound on day 7 after the surgical procedure (Figure 2). The fistula and stenosi...

Watch this Scientific Journal Video about Murine Model of Central Venous Stenosis using Aortocaval Fistula with an Outflow Stenosis at JoVE.com

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

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 ...

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7.3K Views.  Yale University. Centrale veneuze stenose kan zowel aortocaval fistel, of AVF, primaire falen van de rijping, evenals later falen in een werkende fistel veroorzaken. Deze techniek maakt gebruik van een aangepaste versie van een gevestigde murine AVF-model dat de klinische loop van de menselijke AVF samenvat en gemakkelijk onder de knie wordt door middel van verschillende kritieke stappen. Bevestig voor het begin van de procedure een gebrek aan respons op teenknijpen in een negen-tot 11 weken oude C57-Black/6-...

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

Technical Aspects of the Mouse Aortocaval Fistula

Technical aspects of creating an arteriovenous fistula in the mouse are discussed. Under general anesthesia, an abdominal incision is made, and the aorta and inferior vena cava (IVC) are exposed. The proximal infrarenal aorta and the distal aorta are dissected for clamp placement and needle puncture, respectively. Special attention is paid to avoid dissection between the aorta and the IVC. After clamping the aorta, a 25 G needle is used to puncture both walls of the aorta into the IVC. The surrounding connective tissue is used for hemostatic compression. Successful creation of the AVF will show pulsatile arterial blood flow in the IVC. Further confirmation of successful AVF can be achieved by post-operative Doppler ultrasound.

The goal is to produce an arteriovenous fistula that is simple and reproducible. This method does not use sutures or glue adhesive. Therefore the samples can be used with the least amount of foreign materials for analysis.

Technical aspects of creating an arteriovenous fistula in the mouse are discussed. Under general anesthesia, an abdominal incision is made, and the aorta ...

Adult Mouse Venous Hypertension Model: Common Carotid Artery to External Jugular Vein Anastomosis.

The understanding of the pathophysiology of brain arteriovenous malformations and arteriovenous fistulas has improved thanks to animal models. A rat model creating an artificial fistula between the common carotid artery (CCA) and the external jugular vein (EJV) has been widely described and proved technically feasible. This construct provokes a consistent cerebral venous hypertension (CVH), and therefore has helped studying the contribution of venous hypertension to formation, clinical symptoms, and prognosis of brain AVMs and dural AVFs. Equivalent mice models have been only scarcely described and have shown trouble with stenosis of the fistula. An established murine model would allow the study of not only pathophysiology but also potential genetic therapies for these cerebrovascular diseases.
We present a model of arteriovenous fistula that produces a durable intracranial venous hypertension in the mouse. Microsurgical anastomosis of the murine CCA and EJV can be difficult due to diminutive anatomy and frequently result in a non-patent fistula. In this step-by-step protocol we address all the important challenges encountered during this procedure. Avoiding excessive retraction of the vein during the exposure, using 11-0 sutures instead of 10-0, and making a carefully planned end-to-side anastomosis are some of the critical steps. Although this method requires advanced microsurgical skills and a longer learning curve that the equivalent in the rat, it can be consistently developed.
This novel model has been designed to integrate transgenic mouse techniques with a previously well-established experimental system that has proved useful to study brain AVMs and dural AVFs. By opening the possibility of using transgenic mice, a broader spectrum of valid models can be achieved and genetic treatments can also be tested. The experimental construct could also be further adapted to the study of other cerebrovascular diseases related with venous hypertension such as migraine, transient global amnesia, transient monocular blindness, etc.

We describe a method for creating a reliable model of cerebral venous hypertension in the adult mouse. This model has been widely described and tested in the rat. This new counterpart in the mice opens the possibility of using genetic modified animals and thereby broadens the applications of the model.

The understanding of the pathophysiology of brain arteriovenous malformations and arteriovenous fistulas has improved thanks to animal models. A rat model ...

A New Murine Model of Endovascular Aortic Aneurysm Repair

Endovascular aneurysm exclusion is a validated technique to prevent aneurysm rupture. Long-term results highlight technique limitations and new aspects of Abdominal aortic aneurysm (AAA) pathophysiology. There is no abdominal aortic aneurysm endograft exclusion model cheap and reproducible, which would allow deep investigations of AAA before and after treatment. We hereby describe how to induce, and then to exclude with a covered coronary stentgraft an abdominal aortic aneurysm in a rat. The well known elastase induced AAA model was first reported in 19901 in a rat, then described in mice2. Elastin degradation leads to dilation of the aorta with inflammatory infiltration of the abdominal wall and intra luminal thrombus, matching with human AAA. Endovascular exclusion with small covered stentgraft is then performed, excluding any interactions between circulating blood and the aneurysm thrombus. Appropriate exclusion and stentgraft patency is confirmed before euthanasia by an angiography thought the left carotid artery. Partial control of elastase diffusion makes aneurysm shape different for each animal. It is difficult to create an aneurysm, which will allow an appropriate length of aorta below the aneurysm for an easy stentgraft introduction, and with adequate proximal and distal neck to prevent endoleaks. Lots of failure can result to stentgraft introduction which sometimes lead to aorta tear with pain and troubles to stitch it, and endothelial damage with post op aorta thrombosis. Giving aspirin to rats before stentgraft implantation decreases failure rate without major hemorrhage. Clamping time activates neutrophils, endothelium and platelets, and may interfere with biological analysis.

Histological and biochemical modifications after aneurysm endograft exclusion are still unclear. We describe a new model of endograft implantation on a murine aneurysm. Through thrombus analysis with and without persistant circulating blood stream interface, and new endovascular biomaterials evaluation, this model will help to better understand endovascular aneurysm exclusion pathobiology.

Endovascular aneurysm exclusion is a validated technique to prevent aneurysm rupture. Long-term results highlight technique limitations and new aspects of ...

A Novel Murine Model of Arteriovenous Fistula Failure: The Surgical Procedure in Detail

The arteriovenous fistula (AVF) still suffers from a high number of failures caused by insufficient remodeling and intimal hyperplasia from which the exact pathophysiology remains unknown. In order to unravel the pathophysiology a murine model of AVF-failure was developed in which the configuration of the anastomosis resembles the preferred situation in the clinical setting. A model was described in which an AVF is created by connecting the venous end of the branch of the external jugular vein to the side of the common carotid artery using interrupted sutures. At a histological level, we observed progressive stenotic intimal lesions in the venous outflow tract that is also seen in failed human AVFs. Although this procedure can be technically challenging due to the small dimensions of the animal, we were able to achieve a surgical success rate of 97% after sufficient training. The key advantage of a murine model is the availability of transgenic animals. In view of the different proposed mechanisms that are responsible for AVF failure, disabling genes that might play a role in vascular remodeling can help us to unravel the complex pathophysiology of AVF failure.

Here we present a murine model of arteriovenous fistula (AVF) failure in which a clinically relevant anastomotic configuration is incorporated. This model can be used to study the pathophysiology and to test possible therapeutic interventions.

The arteriovenous fistula (AVF) still suffers from a high number of failures caused by insufficient remodeling and intimal hyperplasia from which the ...

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 ...

Immunology and Infection

Murine Cervical Aortic Transplantation Model using a Modified Non-Suture Cuff Technique

With the introduction of powerful immunosuppressive protocols, distinct advances are possible in the prevention and therapy of acute rejection episodes. However, only minor improvement in the long-term results of transplanted solid organs could be observed over the past decades. In this context, chronic allograft vasculopathy (CAV) still represents the leading cause of late organ failure in cardiac, renal and pulmonary transplantation.
Thus far, the underlying pathogenesis of CAV development remains unclear, explaining why effective treatment strategies are presently missing and emphasizing a need for relevant experimental models in order to study the underlying pathophysiology leading to CAV formation. The following protocol describes a murine heterotopic cervical aortic transplantation model using a modified non-suture cuff technique. In this technique, a segment of the thoracic aorta is interpositioned in the right common carotid artery. With the use of the non-suture cuff technique, an easy to learn and reproducible model can be established, minimizing the possible heterogeneity of sutured vascular micro anastomoses.

Here, we present a protocol of heterotopic aortic transplantation in mice using the non-suture cuff technique in a cervical murine model. This model can be used to study the underlying pathology of chronic allograft vasculopathy (CAV) and can help evaluate new therapeutic agents in order to prevent its formation.

With the introduction of powerful immunosuppressive protocols, distinct advances are possible in the prevention and therapy of acute rejection episodes. ...

A Rabbit Venous Interposition Model Mimicking Revascularization Surgery using Vein Grafts to Assess Intimal Hyperplasia under Arterial Blood Pressure

Although vein grafts have been commonly used as autologous grafts in revascularization surgeries for ischemic diseases, the long-term patency remains poor because of the acceleration of intimal hyperplasia due to the exposure to arterial blood pressure. The present protocol is designed for the establishment of experimental venous intimal hyperplasia by interposing rabbit jugular veins to the ipsilateral carotid arteries. The protocol does not require surgical procedures deep in the body trunk and the extent of the incision is limited, which is less invasive for the animals, allowing long-term observation after implantation. This simple procedure enables researchers to investigate strategies to attenuate the progression of intimal hyperplasia of the implanted vein grafts. Using this protocol, we reported the effects transduction of microRNA-145 (miR-145), which is known to control the phenotype of vascular smooth muscle cells (VSMCs) from the proliferative to the contractile state, into harvested vein grafts. We confirmed the attenuation of intimal hyperplasia of vein grafts by transducing miR-145 before implantation surgery through the phenotype change of the VSMCs. Here we report a less invasive experimental platform to investigate the strategies that can be used to attenuate intimal hyperplasia of vein grafts in revascularization surgeries.

The present protocol aims to experimentally create venous intimal hyperplasia by subjecting veins to arterial blood pressure for developing strategies to attenuate venous intimal hyperplasia following revascularization surgery using vein grafts.

Although vein grafts have been commonly used as autologous grafts in revascularization surgeries for ischemic diseases, the long-term patency remains poor ...

A Murine Model of Hemodialysis Access-Related Hand Dysfunction

Chronic kidney disease is a major public health problem, and the prevalence of end-stage renal disease (ESRD) requiring chronic renal replacement therapies such as hemodialysis continues to increase. Autogenous arteriovenous fistula (AVF) placement remains a primary vascular access option for ESRD patients. Unfortunately, approximately half of the hemodialysis patients experience dialysis access-related hand dysfunction (ARHD), ranging from subtle paresthesia to digital gangrene. Notably, the underlying biologic drivers responsible for ARHD are poorly understood, and no adequate animal model exists to elucidate the mechanisms and/or develop novel therapeutics for the prevention/treatment of ARHD. Herein, we describe a new mouse model in which an AVF is created between the left common iliac artery and vein, thereby facilitating the assessment of limb pathophysiology. The microsurgery includes vessel isolation, longitudinal venotomy, creation of arteriovenous anastomosis, and venous reconstruction. Sham surgeries include all the critical steps except for AVF creation. Iliac AVF placement results in clinically relevant alterations in central hemodynamics, peripheral ischemia, and impairments in hindlimb neuromotor performance. This novel preclinical AVF model provides a useful platform that recapitulates common neuromotor perturbations reported by hemodialysis patients, allowing researchers to investigate the mechanisms of ARHD pathophysiology and test potential therapeutics.

This protocol details the surgical steps of murine common iliac arteriovenous fistula creation. We developed this model to study hemodialysis access-related limb pathophysiology.

Chronic kidney disease is a major public health problem, and the prevalence of end-stage renal disease (ESRD) requiring chronic renal replacement ...

Drug Treatment by Central Venous Catheter in a Mouse Model of Angiotensin II Induced Abdominal Aortic Aneurysm and Monitoring by 3D Ultrasound

Since pharmaceutical treatment options are lacking in the clinical management of abdominal aortic aneurysm (AAA), animal models, in particular mouse models, are applied to advance the understanding of the disease pathogenesis and to identify potential therapeutic targets. Testing novel drug candidates to block AAA growth in these models generally requires repeated drug administration during the time course of the experiment. Here, we describe a compiled protocol for AAA induction, insertion of an intravenous catheter to facilitate prolonged therapy, and serial AAA monitoring by 3D ultrasound. Aneurysms are induced in apolipoprotein E (ApoE) deficient mice by angiotensin II release over 28 days from osmotic mini-pumps implanted subcutaneously into the mouse back. Subsequently, the surgical procedure for external jugular vein catheterization is conducted to allow for daily intravenous drug treatment or repeated blood sampling via a subcutaneous vascular access button. Despite the two dorsal implants, the monitoring of AAA development is readily facilitated by sequential semi-automated 3D ultrasound analysis, which yields comprehensive information on the expansion of aortic diameter and volume and on aneurysm morphology, as illustrated by experimental examples.

This protocol describes the consecutive implantation of an osmotic pump to induce abdominal aortic aneurysm by angiotensin II release in apolipoprotein E (ApoE) deficient mice and of a vascular access port with a jugular vein catheter for repeated drug treatment. Monitoring of aneurysm development by 3D ultrasound is effectively conducted despite dorsal implants.

Since pharmaceutical treatment options are lacking in the clinical management of abdominal aortic aneurysm (AAA), animal models, in particular mouse ...

Modified Arteriovenous Fistula Surgery to Establish the RV Volume Overload Mouse Model

Establishment and Confirmation of a Postnatal Right Ventricular Volume Overload Mouse Model

Right ventricular (RV) volume overload (VO) is common in children with congenital heart disease. In view of distinct developmental stages,the RV myocardium may respond differently to VO in children compared to adults. The present study aims to establish a postnatal RV VO model in mice using a modified abdominal arteriovenous fistula. To confirm the creation of VO and the following morphological and hemodynamic changes of the RV, abdominal ultrasound, echocardiography, and histochemical staining were performed for 3 months. As a result, the procedure in postnatal mice showed an acceptable survival and fistula success rate. In VO mice, the RV cavity was enlarged with a thickened free wall, and the stroke volume was increased by about 30%-40% within 2 months after surgery. Thereafter, the RV systolic pressure increased, corresponding pulmonary valve regurgitation was observed, and small pulmonary artery remodeling appeared. In conclusion, modified arteriovenous fistula (AVF) surgery is feasible to establish the RV VO model in postnatal mice. Considering the probability of fistula closure and elevated pulmonary artery resistance, abdominal ultrasound and echocardiography must be performed to confirm the model status before application.

Author Spotlight: Establishment and Confirmation of a Postnatal Right Ventricular Volume Overload Mouse Model  

This protocol presents the establishment and confirmation of a postnatal right ventricular volume overload (VO) model in mice with abdominal arteriovenous fistula (AVF), which can be applied to investigate how VO contributes to postnatal heart development.

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

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