This study was supported by a grant from the Dutch Kidney Foundation (KJPB 08.0003). 
Carolien Rothuizen is acknowledged for her contribution to the study. Hoang Pham is acknowledged for his assistance with the pathology work-up.

All experiments were approved by the committee on animal welfare of the Leiden University Medical Center.
1. Animal Preparation and Anaesthesia
<ol>
	<li>Anesthetize the mouse (1-3 months old) in a continuous anaesthetic induction chamber filled with 3-4% isoflurane.</li>
	<li>Shave the ventral side of the neck and the inside part of the left upper leg using an electrical razor and use a piece of tape to remove the hair.</li>
	<li>Apply ocular ointment on both eyes.</li>
	<li>Position the animal on the heating blanket on which the nose ventilation mask is fixed. Fixate the head to the nose mask using adhesive tape.
	<ol>
		<li>Deliver isoflurane at a concentration of 1.5-2.0% using oxygen enriched air set at the following flow: air 0.3 L/min, 100% oxygen 0.2 L/min. Pinch the skin between the toes to assess the depth of anaesthesia and adjust the concentration of isoflurane if needed.</li>
	</ol>
	</li>
	<li>Fixate the animal to the heating blanket with adhesive tape on all the limbs that is set at approximately 37 &#176;C in a supine position.</li>
	<li>Inject buprenorphine dissolved in sodium chloride (NaCl) 0.9% at a dosage of 0.1 mg/kg together with 0.5 ml&#160;of sterile NaCl 0.9% subcutaneously in the flank of the animal.</li>
	<li>Apply chlorhexidine tincture 0.5% to the prepared area.</li>
</ol>
2. Skin Incision
<ol>
	<li>Position the animal under the microscope with its head towards the surgeon.</li>
	<li>Make a longitudinal incision of approximately 1.5 cm in the midline of the ventral neck using a micro scissor.</li>
	<li>Grab the salivary glands with forceps and displace the right salivary gland cranially until it is partly outside the wound.</li>
</ol>
3. Dissecting and Preparing the Vein

<ol>
	<li>Identify and bluntly dissect the dorsomedial branch of the right external jugular vein completely from its perivascular tissue by spreading just alongside and in the direction of the blood vessel.</li>
	<li>Place a loop (10.0 suture) around the dissected vein and apply one knot without locking it.</li>
</ol>
4. Removing the Sternocleidomastoid Muscle
<ol>
	<li>Using forceps, bluntly dissect the right sternocleidomastoid muscle from its surroundings by spreading along its border. Place a pair of open forceps under the isolated muscle and ligate it proximally and distally with a 6.0 suture. Subsequently, excise the muscle using a cauterizer.</li>
</ol>
5. Dissecting and Preparing the Common Carotid Artery
<ol>
	<li>Identify and dissect the right common carotid artery using forceps. Place a 6.0 suture thread around the artery to aid in the manipulation of the artery.</li>
	<li>Apply vascular clamps as distally and proximally as possible.</li>
	<li>Make a longitudinal incision in the middle of the artery of approximately 1 mm using the specialised micro scissor.</li>
	<li>Rinse the artery with a heparin solution [100 IU/ml] until the vessel is clear of blood.</li>
	<li>Measure and adjust the length of the incision if necessary to match the width the vascular clamp [1.1 mm].</li>
</ol>
6. Ligation of the Vein
<ol>
	<li>Apply a vascular clamp proximally of the already prepared vein and place a suture thread around the vein.</li>
	<li>Apply gentle caudal traction to the vein using a haemostat that is placed on the suture thread and ligate the vein as distally as possible using the 10.0 suture that was placed earlier.</li>
	<li>Cut the vein using a scissor just proximally to the ligation.</li>
	<li>Using vascular forceps, gently open the lumen of the vein and rinse it with a heparin solution [100 IU/ml].</li>
</ol>
7. Creation of Anastomosis Part 1
<ol>
	<li>Connect the vein to the artery with an interrupted suture (10.0) using 2 square knots at the twelve o&#39;clock position followed by a suture at the six o&#39;clock position. Make sure to connect the vein to the artery without allowing it to rotate around its own axis. Rotate the animal in order to improve the surgical exposure.</li>
	<li>Position a suture thread around the vein and apply gentle lateral traction using a haemostat.</li>
	<li>Using the same interrupted sutures, complete the anastomosis by placing approximately 3-4 additional sutures at the visible ventral side of the anastomosis.</li>
</ol>
8. Heparin Administration
<ol>
	<li>Rotate heating blanket 180&#176; and focus the microscope on the left upper leg.</li>
	<li>Identify the femoral vein/artery/nerve bundle by looking for a vascular structure that runs longitudinally in the medial part of the upper leg and can be seen through the skin. Make an incision with the micro scissor of approximately 1 cm directly above the femoral vein in a lengthwise manner.</li>
	<li>Carefully dissect the perivascular tissue of femoral vein and inject heparin at a dose of 0.2 IU/g bodyweight in the femoral vein.</li>
</ol>
9. Creation of Anastomosis Part 2
<ol>
	<li>Return to the neck area and remove the suture thread that was placed around the vein.</li>
	<li>Place a suture thread (6.0) that subsequently passes below the carotid artery, over the vein and again below the carotid artery. (Figure 1L)</li>
	<li>Remove the venous vascular clamp.</li>
	<li>Next, twist the half completed arteriovenous fistula 180&#176; along the axis of the carotid artery in a clockwise manner by simultaneously twisting both the vascular clamps and apply medial traction to the ends of the suture thread.</li>
	<li>Complete the anastomosis in the same fashion as is described in step 7.2.</li>
</ol>
10. Vascular Clamp Removal
<ol>
	<li>Remove the suture thread and rotate the vascular clamps 180&#176; counter clockwise.</li>
	<li>Remove the distal vascular clamp followed by the proximal vascular clamp.</li>
	<li>Assess the patency of the anastomosis by gently occluding the venous outflow tract with the vascular forceps. In case of a patent anastomosis, the pre-occluding venous section will expand in a pulsatile fashion. Relocate the right salivary gland to its original anatomical position using forceps.</li>
</ol>
11. Skin Closure and Postoperative Care
<ol>
	<li>Close the skin of the upper leg with an uninterrupted suture (6.0).</li>
	<li>Close the skin of the neck with an uninterrupted suture (6.0).</li>
	<li>Remove the animal from the heating blanket and inject 0.5 ml NaCl 0.9% subcutaneously.</li>
	<li>Place the animal in a darkened cage that is heated with a heating lamp and let it recover fully. When an animal does not recover fully, make sure it does not suffer from a hemodynamic shock due to bleeding in the surgical area. Look for signs such as a swollen neck and the leakage of blood.</li>
	<li>After approximately 6 hr&#160;after the first injection of buprenorphine, inject a single dose of a sustained-release formulation of buprenorphine subcutaneously in a dose that is recommended by the manufacturer, in order to provide adequate analgesia for an additional 72 hr.&#160; &#160;</li>
</ol>
12. Tissue Harvesting
<ol>
	<li>Anesthetize the animal with an anesthetic-mixture containing midazolam (5 mg/kg), medetomidine (0.5 mg/kg) and fentanyl (0.05 mg/kg) administered intraperitoneally.</li>
	<li>Fixate the animal in a supine position by inserting needles through its paws and cheeks into a non-heated silicon mat.</li>
	<li>Make an incision of approximately 1.5-2.0 cm over the scar in the neck with the micro scissor.</li>
	<li>Dissect the AVF, and place suture threads (6.0) around the proximal-, distal artery and venous outflow tract for easy identification.</li>
	<li>Assess the patency of the AVF as is described in section 10.3 of the protocol.</li>
	<li>Open the peritoneal cavity by performing a medial laparotomy using the micro scissor.
	<ol>
		<li>Cut through the left and right ventral rib cage using a pair of scissors starting caudally in the midclavicular line. Using a micro scissor, transect the diaphragm that is attached to the middle part of the ventral rib cage and subsequently displace this section cranially using forceps.</li>
		<li>Locate the inferior caval vein and transect it with a micro scissor.</li>
	</ol>
	</li>
	<li>Insert a needle 23 gauge needle in the left ventricle and perfuse with PBS at a pressure of approximately 100 mmHg until the intravasal fluid is nearly clear.</li>
	<li>Without removing the needle, now perfuse with 4% formalin for 10 min at a pressure of approximately 100 mmHg.</li>
	<li>Excise the AVF by transecting both the arteries and vein with the micro scissor and submerge it in a 4% formalin solution O/N.</li>
</ol>
13. Tissue Embedding and Sectioning
<ol>
	<li>Process the tissue to paraffin according to the manufacture&#39;s protocol. The protocol applied consisted of the following:
	<ol>
		<li>Immerse in Ethanol 70% for 1 hr at RT. Repeat 2x.</li>
		<li>Immerse in Ethanol 96% for 1 hr at RT. Repeat 2x.</li>
		<li>Immerse in Ethanol 99.5% for 1 hr at RT. Repeat once.</li>
		<li>Immerse in Ethanol 99.5% for 1.5 hr at RT.</li>
		<li>Immerse in xylene 100% for 1 hr at RT. Repeat once.</li>
		<li>Immerse in Parafin for 1 hr at 62 &#176;C. Repeat 2x.</li>
	</ol>
	</li>
	<li>Embed the AVF in paraffin so that the venous outflow tract is orientated perpendicular to the embedding cassette.</li>
	<li>Using a microtome, create 12 serial sections each consisting of 30 sections with a 5 &#181;m thickness. Then of each serial section, place one section on a single mounting glass starting from position one up to twelve in an orderly fashion starting with the area closest to the anastomosis.</li>
</ol>

<ol>
	<li>Rotmans, J. I. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Animal+Models+for+Studying+Pathophysiology+of+Hemodialysis+Access.">Animal Models for Studying Pathophysiology of Hemodialysis Access.</a> The Open Urology &amp; Nephrology Journal. 7, 14-21 (2014).</li><li>Kwei, 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=Early+adaptive+responses+of+the+vascular+wall+during+venous+arterialization+in+mice.">Early adaptive responses of the vascular wall during venous arterialization in mice.</a> Am.J.Pathol. 164 (1), 81-89 (2004).</li><li>Castier, Y., 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=Characterization+of+neointima+lesions+associated+with+arteriovenous+fistulas+in+a+mouse+model.">Characterization of neointima lesions associated with arteriovenous fistulas in a mouse model.</a> Kidney Int. 70 (2), 315-320 (2006).</li><li>Krishnamoorthy, M. 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=Hemodynamic+wall+shear+stress+profiles+influence+the+magnitude+and+pattern+of+stenosis+in+a+pig+AV+fistula.">Hemodynamic wall shear stress profiles influence the magnitude and pattern of stenosis in a pig AV fistula.</a> Kidney Int. 74 (11), 1410-1419 (2008).</li><li>Ene-Iordache, B., Cattaneo, L., Dubini, G., 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=Effect+of+anastomosis+angle+on+the+localization+of+disturbed+flow+in+'side-to-end'+fistulae+for+haemodialysis+access.">Effect of anastomosis angle on the localization of disturbed flow in 'side-to-end' fistulae for haemodialysis access.</a> Nephrol. Dial. Transplant. 28 (4), 997-1005 (2013).</li><li>Wong, C. Y., 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=Vascular+remodeling+and+intimal+hyperplasia+in+a+novel+murine+model+of+arteriovenous+fistula+failure.">Vascular remodeling and intimal hyperplasia in a novel murine model of arteriovenous fistula failure.</a> J.Vasc.Surg. 59 (1), 192-201 (2014).</li><li>Rekhter, M., Nicholls, S., Ferguson, M., Gordon, D. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Cell+proliferation+in+human+arteriovenous+fistulas+used+for+hemodialysis.">Cell proliferation in human arteriovenous fistulas used for hemodialysis.</a> Arterioscler. Thromb. 13 (4), 609-617 (1993).</li><li>Falk, A. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Maintenance+and+salvage+of+arteriovenous+fistulas.">Maintenance and salvage of arteriovenous fistulas.</a> J. Vasc. Interv. Radiol. 17 (5), 807-813 (2006).</li><li>Tordoir, J. H., 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=Prospective+evaluation+of+failure+modes+in+autogenous+radiocephalic+wrist+access+for+haemodialysis.">Prospective evaluation of failure modes in autogenous radiocephalic wrist access for haemodialysis.</a> Nephrol. Dial. Transplant. 18 (2), 378-383 (2003).</li><li>Dixon, B. S., Novak, L., Fangman, J. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Hemodialysis+vascular+access+survival:+upper-arm+native+arteriovenous+fistula.">Hemodialysis vascular access survival: upper-arm native arteriovenous fistula.</a> Am. J. Kidney Dis. 39 (1), 92-101 (2002).</li><li>Yang, B., Shergill, U., Fu, A. A., Knudsen, B., Misra, S. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=The+mouse+arteriovenous+fistula+model.">The mouse arteriovenous fistula model.</a> J. Vasc. Interv. Radiol. 20 (7), 946-950 (2009).</li><li>Kang, 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=Regional+and+systemic+hemodynamic+responses+following+the+creation+of+a+murine+arteriovenous+fistula.">Regional and systemic hemodynamic responses following the creation of a murine arteriovenous fistula.</a> Am. J. Physiol Renal Physiol. 301 (4), F845-F851 (2011).</li><li>Wong, C. Y., 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=Elastin+is+a+key+regulator+of+outward+remodeling+in+arteriovenous+fistulas.">Elastin is a key regulator of outward remodeling in arteriovenous fistulas.</a> Eur. J. Vasc. Endovasc. Surg. 49 (4), 480-486 (2015).</li><li>Kennedy, R., 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=Does+renal+failure+cause+an+atherosclerotic+milieu+in+patients+with+end-stage+renal+disease.">Does renal failure cause an atherosclerotic milieu in patients with end-stage renal disease.</a> Am. J. Med. 110 (3), 198-204 (2001).</li><li>Cheung, A. 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=Atherosclerotic+cardiovascular+disease+risks+in+chronic+hemodialysis+patients.">Atherosclerotic cardiovascular disease risks in chronic hemodialysis patients.</a> Kidney Int. 58 (1), 353-362 (2000).</li><li>Lee, T., 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=Severe+venous+neointimal+hyperplasia+prior+to+dialysis+access+surgery.">Severe venous neointimal hyperplasia prior to dialysis access surgery.</a> Nephrol. Dial. Transplant. 26 (7), 2264-2270 (2011).</li><li>Kokubo, T., 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=CKD+accelerates+development+of+neointimal+hyperplasia+in+arteriovenous+fistulas.">CKD accelerates development of neointimal hyperplasia in arteriovenous fistulas.</a> J. Am. Soc. Nephrol. 20 (6), 1236-1245 (2009).</li></ol>

The authors have nothing to disclose.&#160;

The AVF is considered to be the Achilles&#39; heel in hemodialysis treatment. Unfortunately, the AVF still suffers from a high number of failure8-10. Despite extensive research on the underlying mechanisms, the exact pathophysiology remains unknown. Numerous murine models for AVF failure have already been described in literature2,3,11,12. However, none of these models incorporate a venous end to arterial side anastomosis configuration that is most used in the clinical situation. This is very relevan...

A functional vascular access conduit is of vital importance for patients with renal failure that depend on chronic hemodialysis to stay alive. The construction of an arteriovenous fistula (AVF) is currently the preferred choice for vascular access. However, AVF related complications constitute a major cause of morbidity for patients on chronic hemodialysis. Despite extensive scientific efforts, none of the novel approaches to reduce AVF access-related complications did result in substantial improvement of AVF durability. Part of this disappointing progress relates to incomplete understanding of the underlying pathophysiology of hemodialysis access failure.
To unravel the pathophysiology of AV access failure, animal models that closely mimic human pathology are of utmost importance. In this respect, not only the animal species but also the anastomotic site, the required anti-coagulatory therapy and the duration of follow up after surgery should be taken into account1. While large animals are the most suitable for intervention studies aimed to develop new therapeutic strategies, murine models have the greatest potential to gain more insight in the molecular mechanisms underlying AV access failure due to the availability of transgenic mice. In addition, a large number of mice can be used for this purpose at lower costs compared to larger animal use.
The first murine model of AVF failure was described in 2004 by Kwei and et al.2 In this model, AVFs were constructed using the carotid artery and the jugular vein in an end-to-end manner using an intravascular catheter. This model could be useful to study early venous adaptation in AVFs although the end-to-end configuration and the presence of an intravascular catheter limit the validity of this model for human AVFs. An improved AVF model was introduced by Castier and et al.3 in which the end of the carotid artery is connected to the side of the jugular vein. However, AVFs in hemodialysis patients are usually constructed by anatomizing the end of a vein to the side of an artery. The exact configuration of the AVF is a crucial characteristic of an AV access model since it determines the hemodynamic profile within the conduit4. The latter is an important contributor to endothelial dysfunction and subsequent development of intimal hyperplasia (IH)5.
A novel murine model was recently developed with an identical anatomical configuration as is utilized in humans6. In this model, AVF are created in C57BL/6 mice by anastomosing the end of a branch of the external jugular vein to the side of the common carotid artery with interrupted sutures. In the present paper, we focus on the microsurgical procedure of this model in order to facilitate the widespread use of this murine model, aimed to unravel the complex pathophysiology of hemodialysis access failure.

<table><tbody><tr><td>Dissecting microsocpe</td><td>Leica</td><td>M80</td><td></td></tr><tr><td>Forceps</td><td>Medicon</td><td>07.61.25</td><td></td></tr><tr><td>Vascular forceps</td><td>S&amp;amp;T</td><td>JFL-3D.2</td><td></td></tr><tr><td>Vascular forceps</td><td>S&amp;amp;T</td><td>D-5a.2</td><td></td></tr><tr><td>Forceps</td><td>Roboz</td><td>SS/45</td><td></td></tr><tr><td>Micro scissor 5 mm blade</td><td>Fine science tools</td><td>15000-08</td><td></td></tr><tr><td>Micro scissor 2 mm blade</td><td>Fine science tools</td><td>15000-03</td><td></td></tr><tr><td>Scissor</td><td>Medicon</td><td>05.12.21</td><td></td></tr><tr><td>Clip applier 1</td><td>S&amp;amp;T</td><td>CAF-4</td><td></td></tr><tr><td>Vascular clamp 1</td><td>S&amp;amp;T</td><td>B-1V</td><td></td></tr><tr><td>Clip applier 2</td><td>BBraun</td><td>FE572K</td><td></td></tr><tr><td>Vascular clamp 2</td><td>BBraun</td><td>FE740K</td><td></td></tr><tr><td>Hemostatic forceps</td><td>BBraun</td><td>BH110</td><td></td></tr><tr><td>10.0 sutures</td><td>BBraun</td><td>G1117041</td><td></td></tr><tr><td>6.0 sutures</td><td>BBraun</td><td>768464</td><td></td></tr><tr><td>Cauterizer</td><td>Fine science tools</td><td>18010-00</td><td></td></tr><tr><td>Needle holder</td><td>Medicon</td><td>11.82.18</td><td></td></tr><tr><td>Ocular ointment</td><td>Pharmachemie</td><td>41821101</td><td></td></tr><tr><td>Chlorhexidine tincture 0,5%</td><td>Leiden University Medical Center</td><td>NA</td><td></td></tr><tr><td>Heparin</td><td>Leo Pharma</td><td>012866-08</td><td></td></tr><tr><td>Buprenorphin</td><td>RB Pharmaceuticals&amp;nbsp;</td><td>283732</td><td></td></tr><tr><td>Isoflurane</td><td>Pharmachemie</td><td>45,112,110</td><td></td></tr><tr><td>Anesthesia mask</td><td>Maastricht university</td><td>custom made</td><td></td></tr><tr><td>Midazolam</td><td>Actavis</td><td>AAAC6877</td><td></td></tr><tr><td>Dexmedetomidine</td><td>Orion</td><td>141-267</td><td></td></tr><tr><td>Fentanyl</td><td>Bipharma</td><td>15923002</td><td></td></tr><tr><td>Continuous anaesthetic induction chamber</td><td>Vet-tech solutions</td><td>AN010R</td><td></td></tr></tbody></table>

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.

After the creation of the anastomosis (Figure 1), the patency should be assessed by shortly occluding the venous outflow tract with a vascular forceps. When the anastomosis is patent, the vascular tract proximal to the occlusion should clearly expand in a pulsatile manner. In addition, the patency is confirmed by using near infrared fluoroscopy (NIRF) that effectively functions as an angiography (Figure 2). A failure in the surgical procedure can lead to an occlusion of the anastomosis a...

Watch this Scientific Journal Video about A Novel Murine Model of Arteriovenous Fistula Failure: The Surgical Procedure in Detail at JoVE.com

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

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 ...

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12.9K Views.  Leiden University Medical Center. 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.

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

Microsurgical Venous Pouch Arterial-Bifurcation Aneurysms in the Rabbit Model: Technical Aspects

For ruptured human cerebral aneurysms endovascular embolization has become an equivalent alternative to aneurysm clipping.1 However, large clinical trials have shown disappointing long-term results with unacceptable high rates of aneurysm recanalization and delayed aneurysm rupture.2 To overcome these problems, animal experimental studies are crucial for the development of better endovascular devices.3-5
Several animal models in rats, rabbits, canines and swine are available.6-8 Comparisons of the different animal models showed the superiority of the rabbit model with regard to hemodynamics and comparability of the coagulation system and cost-effectiveness.9-11 
The venous pouch arterial bifurcation model in rabbits is formed by a venous pouch sutured into an artificially created true bifurcation of both common carotid arteries (CCA). The main advantage of this model are true bifurcational hemodynamics.12 The major drawbacks are the sofar high microsurgical technical demands and high morbidity and mortality rates of up to 50%.13 These limitations have resulted in less frequent use of this aneurysm model in the recent years. These shortcomings could be overcome with improved surgical procedures and modified peri- and postoperative analgetic management and anticoagulation.14-16 Our techniques reported in this paper demonstrate this optimized technique for microsurgical creation of arterial bifurcation aneurysms.

An optimized technique for the microsurgical creation of arterial bifurcation aneurysms mimicking bifurcation human cerebral aneurysms is described. A venous pouch is sutured into an artificially created true bifurcation of both common carotid arteries. Facilitated microsurgical techniques and aggressive postoperative anticoagulation and analgesia lead to minimized morbidity rates and high aneurysm patency rates.

For ruptured human cerebral aneurysms endovascular embolization has become an equivalent alternative to aneurysm clipping.1 However, large clinical trials ...

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 ...

Murine Model of Femoral Artery Wire Injury with Implantation of a Perivascular Drug Delivery Patch

Percutaneous interventions including balloon angioplasty and stenting have been used to restore blood flow in vessels with occlusive vascular disease. While these therapies lead to the rapid restoration of blood flow, these technologies remain limited by restenosis in the case of bare metal stents and angioplasty, or reduced healing and possibly enhanced risk of thrombosis in the case of drug eluting stents. A key pathophysiological mechanism in the formation of restenosis is intimal hyperplasia caused by the activation of vascular smooth muscle cells and inflammation due to arterial stretch and injury. Surgeries that induce arterial injury in genetically modified mice are useful for the mechanistic study of the vascular response to injury but are often technically challenging to perform in mouse models due to the their small size and lack of appropriate sized devices. We describe two approaches for a surgical technique that induces endothelial denudation and arterial stretch in the femoral artery of mice to produce robust neointimal hyperplasia. The first approach creates an arteriotomy in the muscular branch of the femoral artery to obtain vascular access. Following wire injury this arterial branch is ligated to close the arteriotomy. A second approach creates an arteriotomy in the main femoral artery that is later closed through localized cautery. This method allows for vascular access through a larger vessel and, consequently, provides a less technically demanding procedure that can be used in smaller mice. Following either method of arterial injury, a degradable drug delivery patch can be placed over or around the injured artery to deliver therapeutic agents.

We describe a surgical technique that produces wire injury in the femoral artery of mice to induce neointimal hyperplasia to serve as a model testing system for the perivascular delivery of therapeutic compounds for the inhibition of restenosis.

Percutaneous interventions including balloon angioplasty and stenting have been used to restore blood flow in vessels with occlusive vascular disease. ...

Intraluminal Drug Delivery to the Mouse Arteriovenous Fistula Endothelium

Delivery of therapeutic agents to enhance arteriovenous fistula (AVF) maturation can be administered either via intraluminal or external routes. The simple murine AVF model was combined with intraluminal administration of drug solution to the venous endothelium at the same time as fistula creation. Technical aspects of this model are discussed. Under general anesthesia, an abdominal incision is made and the aorta and inferior vena cava (IVC) are exposed. The infra-renal aorta and IVC are dissected for clamping. After proximal and distal clamping, the puncture site is exposed and a 25 G needle is used to puncture both walls of the aorta and into the IVC. Immediately after the puncture, a reporter gene-expressing viral vector was infused in the IVC via the same needle, followed by 15 min of incubation. The intraluminal administration method enabled more robust viral gene delivery to the venous endothelium compared to administration by the external route. This novel method of delivery will facilitate studies that explore the role of the endothelium in AVF maturation and enable intraluminal drug delivery at the time of surgical operation.

After puncturing the aorta through the inferior vena cava (IVC) to create an aorto-caval fistula in the mouse, solution containing a drug is infused into the IVC via the same needle, followed by incubation. This method enables more robust drug delivery to the venous endothelium compared to the external route.

Delivery of therapeutic agents to enhance arteriovenous fistula (AVF) maturation can be administered either via intraluminal or external routes. The ...

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 ...

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 ...

A Modified Technique for Arteriovenous Fistula Construction in Rabbits

Juxta-anastomotic stenosis is a challenging problem that often causes non-maturation and decreases the patency of an arteriovenous fistula (AVF). Injury to the veins and arteries during the operation and hemodynamic changes can lead to intimal hyperplasia, leading to juxta-anastomotic stenosis. To reduce injury to the veins and arteries during the operation, this study proposes a new modified no-touch technique (MNTT) for AVF construction that can decrease the rate of juxta-anastomotic stenosis and improve the AVF patency. To unravel the hemodynamic changes and mechanisms of the MNTT, this study presented an AVF procedure using this technique. Although this procedure is technically challenging, 94.4% procedural success was achieved after adequate training. Ultimately, 13 out of 34 rabbits had a functional AVF 4 weeks after the surgery, leading to a 38.2% AVF patency rate. However, at 4 weeks, the survival rate was 86.1%. Ultrasonography showed active blood flow through AVF anastomosis. Furthermore, the spiral laminar flow was observed in the vein and artery near the anastomosis, suggesting that this technique may improve the hemodynamics of the AVF. On histological observation, significant venous intimal hyperplasia was observed at the AVF anastomosis, whereas no significant intimal hyperplasia was observed at the proximal external jugular vein (EJV) of the anastomosis. This technique will improve the understanding of the mechanisms underlying the use of MNTT for AVF construction and provide technical support for the further optimization of the surgical approach in AVF construction.

The present protocol proposes the creation of an arteriovenous fistula in rabbits using a modified no-touch technique. The technique involves the side-to-side anastomosis of the common carotid artery and external jugular vein without the dissection of the perivenous tissues or cutting off the artery.

Juxta-anastomotic stenosis is a challenging problem that often causes non-maturation and decreases the patency of an arteriovenous fistula (AVF). Injury ...

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.