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

Summary

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.

Abstract

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.

Introduction

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 mature^1,2,3; 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 maturation^5,6. 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 catheters^7,8.

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.

Protocol

All experiments were performed with approval from the Yale University Institutional Animal Care and Use Committee (IACUC).

1. Anesthesia and pre-operative procedures

1. Sterilize all surgical instruments and materials by autoclaving. Turn on the thermal support device to be certain it is warm (40–42 °C).
2. Place a 9–11-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.
3. 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.
4. Remove fur from the ventral side of the neck to the lower abdomen using Nair, a hair remover.
5. Cleanse and disinfect the surgical site by using a two-step scrub with 10% povidone-iodine and 70% isopropanol. Apply a surgical drape.

2. Operative procedures

1. Exposure of the clamp and puncture sites
   1. Prepare sterile instruments and wear sterile gloves to maintain sterility throughout the surgery.
   2. 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.
   3. 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.
   4. 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.
   5. 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.
2. IVC dissection
   1. 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.
   2. 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.
3. AVF creation
   1. Bend a 25 G needle to a 45–60° angle at a point ~4 mm from the needle tip.
   2. Clamp the infra-renal aorta and IVC by applying a microsurgical clip.
   3. 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.
   4. Holding the aorta in a suitable position, puncture through the aorta into the IVC using the prepared 25 G needle (Figure 1C).
   5. 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.
4. Creation of the IVC stenosis
   1. 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.
   2. 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.
   3. Return organs to their original positions and close the abdomen with 6-0 sutures.

3. Post-operative procedures

1. 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 h following the surgical procedures and thereafter as needed.
2. Confirm the AVF patency postoperatively by using Doppler ultrasound (see the Table of Materials). In addition, measure other vessel and flow characteristics as required.

Results

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...

Discussion

The murine AVF model has been used to study the basic mechanisms and molecular events leading to AVF maturation^13,14. 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 ...

Materials

List of materials used in this article

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